5780 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

Cite this: Chem. Soc. Rev., 2012, 41, 5780–5799

Nanomaterials in complex biological systems: insights from

Raman spectroscopy

Daniela Drescherab

and Janina Kneipp*ab

Received 5th April 2012

DOI: 10.1039/c2cs35127g

The interaction of nanomaterials with biomolecules, cells, and organisms plays an important role

in cell biology, toxicology, and nanotechnology. Spontaneous Raman scattering can be used to

probe biomolecules, cells, whole animals, and nanomaterials alike, opening interesting avenues to

study the interaction of nanoparticles with complex biological systems. In this review we discuss

work in biomedical Raman spectroscopy that has either been concerned directly with

nanostructures and biosystems, or that indicates important directions for successful future studies

on processes associated with nano-bio-interactions.

1 Introduction

The interaction of nanomaterials with biological objects,

i.e., with biomolecules, cells, tissues is of interest in several

different contexts: the awareness of the omnipresence of

nanoparticles in our everyday life has increased tremendously

over the last decade, and has been pushing research in the field

of nanotoxicity. At the same time, significant progress has

been made in nanomaterials research, nanobiotechnology,

and nanophotonics, including work on nanoobjects which

are relevant for the interaction of biological samples with

man-made nanomaterials ‘‘in the field’’. The schematic in

Fig. 1 illustrates some of the questions that are asked when

a nanomaterial encounters a biological cell. The material may

enter the cell, depending on its properties. Will it enter the cell?

In case it does, what is the mechanism of uptake? Where in

the cell will it reside? Will its presence lead to changes in the

cellular biochemistry? If so, what are these changes? Can the

material be considered cytotoxic?

Raman probing and imaging of tissues and cells has become an

established area of research in connection with biodiagnostics and

with the biochemical characterization of complex biomaterials. As

numerous examples have shown, Raman microspectra can be

used for hyperspectral imaging, classification, and identification of

biological samples.1–4 At the same time, they can provide informa-

tion about the structure, and also dynamics of biomolecules.5–8

aHumboldt-Universitat zu Berlin, Department of Chemistry,Brook-Taylor-Str. 2, 12489 Berlin, Germany.E-mail: Janina.kneipp@chemie.hu-berlin.de;Fax: +49-30-2093 6985; Tel: +49-30-2093 7171

b BAM Federal Institute for Materials Research and Testing,12489 Berlin, Germany

Daniela Drescher

Daniela Drescher studiedchemistry at the Humboldt-Universitat zu Berlin, whereshe received her Diplomadegree in 2008. She iscurrently working on herdoctoral thesis in the group ofJanina Kneipp, focusing on theapplication of spectroscopicand microscopic methods toinvestigate the impact ofnanomaterials on eukaryoticcells.

Janina Kneipp

Janina Kneipp received herdoctoral degree from FreieUniversitat Berlin in 2002.After postdoctoral researchat Erasmus UniversiteitRotterdam and PrincetonUniversity’s Chemistry Depart-ment, she has been workingat BAM Federal Institute forMaterials Research and Testingand at the Department ofChemistry of Humboldt-Universitat, where she wasappointed as assistant profes-sor in 2008 and full professorin 2012. Her research interests

lie in the area of vibrational spectroscopies and microscopies toinvestigate small-scale complex biomaterials.

Chem Soc Rev Dynamic Article Links

www.rsc.org/csr CRITICAL REVIEW

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online / Journal Homepage / Table of Contents for this issue

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5781

In the past seven to ten years, several papers have been

published that have as a subject Raman spectroscopy and imaging

of cells, with nanomaterials serving different purposes, ranging

from the application of new labels in imaging approaches, to the

investigation of cellular physiology. In many cases, nanomaterials

are used to generate optical signals in Raman biomedical applica-

tions, such as single-walled carbon nanotubes (SWCNTs)9,10 and

SERS labels.11,12 In others, the properties of vehicles for drug

delivery or photothermal therapy, such as polymeric, SWCNT,

and gold nanocarriers, are studied.13–15

In several recent examples, nanomaterials have been used to

improve Raman signals of the biological samples themselves,

using surface enhanced Raman scattering (SERS) in the local

optical fields of gold or silver nanoparticles.16,17 Due to the

confinement of the local fields to the immediate proximity of

the nanoparticles, the excitation and scattering volumes are

drastically reduced compared to the actual resolution of the

optical microscope. Thereby, only molecules that reside in the

nanometer-scaled vicinity of the nanoparticles contribute to

the observed spectrum, even if the focal volume defined by the

excitation laser and geometry of the setup is much larger.

Molecules can be introduced with the nanoparticles into cells

and organisms, and the nanoparticles can be traced as labels.

Nanomaterials may also provoke a known cellular reaction

that could then be mapped by Raman imaging, such as the

activation of specific enzymes in phagocytosis, thereby enabling

investigations of such processes.18 Also examples of experiments

can be found where Raman spectroscopic characterization and

classification of cells have been undertaken in order to find out

about the toxicity of the nanomaterial.19,20

Spontaneous Raman scattering microscopy, in non-resonant,

resonant, and surface-enhanced experiments, is extremely versatile

regarding the many alterations in the spectroscopic fingerprints of

cells that can be observed, also over time.4,21–24 Furthermore,

micro-Raman experiments enable the combination of the spectro-

scopic experiments with optical manipulation tools, e.g. optical

tweezers to trap cells, their organelles, or the nanomaterials.25–30

Themolecular information in the Raman spectroscopic fingerprints

is connected with a multitude of cellular parameters. In addition to

providing phenotypic data, that can be used to classify and identify

different cells or tissue types and to discriminate them based on

pattern recognition, the Raman spectra of cells reveal information

about molecular structure, interactions and compositional changes.

Regarding this aspect, Raman microscopy is much different from

other microscopic methods that are used to study nanomaterials in

cells, and that provide mainly morphological information along

with signals from the nanomaterial, such as transmission electron

microscopy, dark field microscopy, or fluorescence microscopy.

Using Raman spectroscopy, cellular parameters that are of

importance to the process of the nano-bio-interaction and to

its consequences for the cellular biochemistry may be identified.

Both parts of the nano-bio-system, the molecules of the cell and

those of the nanomaterial, can be studied.

In the context of this article, Raman spectroscopy provides

different perspectives on the nano-bio-systems. Raman scattering

has become an important tool for analyzing the composition of a

complex mixture of biomolecules such as a cell or even a whole

organism. Further, as a basic tool for biophysical structure

analysis, normal and resonant Raman spectroscopy have been

used for decades to find out about the structure and dynamics of

biomolecules. Therefore, even though the information that is

generated in a Raman spectroscopic experiment is always based

on molecular vibrations, different Raman experiments deliver

information about a biological system at very different scales

regarding molecular and morphological complexity and physical

size. First, the macroscopic scale, represented by imaging

applications in whole animals and assessed through innovative

technical and data analysis approaches, will enable us to

better understand systemic influences of nanomaterials on the

organism. Second, at the microscopic scale, Raman images of

individual cells in culture or in tissues, and Raman spectra from

cellular organelles can improve our insight into alterations

occurring in the biochemical composition or interactions in

complex molecular mixtures. Thereby we may gain insight into

the processes that are associated with the response of individual

cells to nanoparticles. Third, the interaction of the nanomaterial

with cellular molecules takes place at the nanoscale. To a great

extent, it involves Raman studies on the structure and structural

changes of purified or isolated molecular species and model

systems of interest, which cannot be covered in-depth here. Of

this third aspect, we will include here normal Raman and SERS

spectra, providing information on the surface interaction of the

nanomaterials themselves inside the biological system.

We will discuss here Raman studies of eukaryotic (that is,

nucleus containing) cells mainly of animal origin under the aspect,

which type of the information can be retrieved from their Raman

spectra during interaction with nanomaterials. We feel that such a

discussion may help to design and perform decisive Raman experi-

ments that employ the great potential of Raman spectroscopy for

studies on the influence of nanoparticles on cells and organisms.

2 Nanomaterials in the organism

Before nanomaterials, targeted or un-wanted, can enter a cell,

they have to come into its proximity. In an animal, this means

Fig. 1 Motivation (schematic). Aspects that are of interest in the

interaction of nanoparticles with cells, with emphasis on potential

nanotoxicity. Toxicity of the nanomaterial may strongly depend on

the particle properties, and involves many biochemical parameters.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5782 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

that the material must enter the organism. Unless the cell

under discussion is an epithelial cell where a nanoparticle

would have direct access, it will encounter different tissues in

the body of the animal, including the blood, or the interstitial

fluid. In many cases, the surface properties and size of the

nanomaterial, the molecules in the body fluids, such as serum

components, and the cell types the material can interact with

will determine its fate in the organism. In this section, we

discuss work on Raman scattering from whole organisms and

ex vivo tissues. The number of Raman experiments that have

been reported with nanoparticles and tissues of animal origin

exceed by far those dealing with plants, probably since the

need for improved tumor diagnostics, imaging and therapeutic

tools has pushed this field of research. Nevertheless, we will

spend a moment summarizing the main directions of Raman

microscopy experiments in plant tissues, before we continue to

review work reported in animals.

2.1 Raman experiments on plant cells

Raman microscopy has become an important tool to study

plant cells as well. As an example, the characterization of plant

cell walls by Raman spectroscopy dates back to the first days

of Raman biospectroscopy.31,32 Cellulose, and also pectin and

lignin have been in the focus of most of the more recent

Raman imaging studies,33–35 in addition to work on other

compounds such as coniferin36 and arabinoxylan.37 Normal

and resonant Raman spectroscopic characterization of the

lipid, chlorophyll and carotenoid building blocks of micro-

algae could have potential use in biotechnology in the future.38–41

Recently, we utilized Raman microspectroscopy for the charac-

terization of molecular and metabolic parameters during the

growth of the pollen tube under varying nutrient conditions

in vitro.42 The results provided evidence of changes in the

chemical composition of the cells during germination and

species-specific utilization of metabolite storage. Nevertheless,

only very few papers report on SERS in plant materials.43–47

A better understanding of the nanomaterial–plant interaction

would be beneficial, particularly because of issues concerned

with nanotoxicity and other physiological aspects. As indicated

in a recent Raman study in tomato plants using conjugates of

SWCNTs and semiconductor quantum dots,48 Raman scattering

may be very useful in the future to elucidate these processes, even

if most of the work reported so far was done with cells of animal

origin. Although many things concerned with nanoparticle

uptake may work entirely differently in plant cells, these cells

share several biochemical aspects with the cells of animal tissues.

Similarly, several Raman spectroscopic investigations have been

carried out on yeast cells that can provide valuable information

about their physiological status, such as viability and cell wall

composition.51–55

2.2 Resonant Raman scattering from carbon nanotubes in

animals

Single-walled carbon nanotubes (SWCNTs) are very promising

tools for bioimaging,10 photothermal therapy, and delivery of

molecular cargo into cells and organs.56,57 By exploiting the

characteristic resonant Raman spectrum of SWCNTs,58 it is

possible to follow the pathway of this nanomaterial in the

body of an animal. When SWCNTs were functionalized with

phospholipids bearing polyethylene glycols (PEG) linked to

an arginine–glycine–apartic acid (RGD) peptide, they spent

a long time in the blood circulation, and were efficiently

accumulated in tumor tissue.14 Evidence of this was provided by

the Raman spectrum of homogenized and dried tissue from the

tumor, liver, kidney and muscle of mice who had received an

intravenous injection of the nanotubes, revealing the characteristic

G band at 1580 cm�1 and the 230 cm�1 A1g radial breathing mode

of the SWCNTs.14 Similarly, Raman spectra of homogenates of the

liver, lung, and spleen tissue have supported toxicological profiling

and could be used to determine the long-term accumulation of

SWCNTs in some organs to even 90 days post-injection of

SWCNTs into the tail vein of mice.59 Interestingly, even though

the SWCNTs were retained for such a long time in the body of the

animals, toxicity in this specific study was relatively low, as was

indicated by biochemical parameters in the serum and other

biological toxicity tests.59 In another toxicology study, the Raman

microscopic characterization of paraffin-embedded, glass mounted

sections of the liver and spleen tissue revealed that non-covalently

PEG-functionalized SWCNTs, as well as oxidized, covalently

functionalized SWCNTs accumulate in both organs, preferably in

the liver, where they were found to persist in Kupffer cells (liver

macrophage cells) for four months.60 Also there, PEG functiona-

lized SWCNTs were not toxic to the animals.60 In vitro toxicity

work by Zhang et al. using the neuronal cell line PC12 demon-

strated that SWCNTs show concentration-dependent and surface

coating-dependent cytotoxicity, with PEG-functionalized SWCNTs

being less cytotoxic than uncoated SWCNTs.61

A major advantage of Raman spectroscopy for imaging and

characterization of biological tissues lies in the possibility to use

wavelengths in the NIR for excitation. The long penetration depth

and low autofluorescence in this spectral region allow for Raman

imaging of whole small animals. Zavaleta et al. reported imaging

of SWCNTs in tumors in nude mice at a depth of 2 mm in the

animal, with raster scan images of the SWCNT G-band intensity,

taken at a step size of 750 mm and 3 s accumulation per

spectrum.12,62 Thereby, small tumors were screened in reasonable

amounts of time. The data from this in vivoRaman imaging study

confirmed the above-cited work of Liu et al. on tissue homo-

genates, who had shown that RGD-functionalized SWCNTs were

targeted to integrin-positive tumor tissues more efficiently than

plain SWCNTs.14 In vivo imaging enables the observation of the

same animal over time. While whole Raman images of organs and

tumors can be taken at specific time points after administration of

SWCNTs,62 time-resolved Raman spectroscopy at the millisecond

time scale can be used for real-time detection of carbon nanotubes

that circulate in living animals.63 After intravenously administering

these nanostructures in the tail vein of the rat, their circulation was

observed by resonant Raman scattering in blood microvessels of

an intact rat ear.63 In ex vivo studies on tissues of mice, Raman

spectroscopy detected SWCNTs in the intestine, feces, kidney,

and bladder, suggesting excretion and clearance of the SWCNTs

via the biliary and renal pathways.64

2.3 SERS tags in living animals

Similar to the detection of SWCNTs, also other strong Raman

scatterers can be observed by in vivo Raman spectroscopy in

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5783

the bodies of live animals. SERS nanoparticle tags, that is,

metal nanostructures with adsorbed molecules providing

a characteristic SERS spectrum, and encapsulated with a

polymer shell, can be introduced into animals and targeted

to tumors as well. As shown by Qian et al., who injected

subcutaneously and intramuscularly in mice SERS tags

consisting of gold nanoparticles bearing malachite green as a

reporter molecule and thiol-modified PEG as a stabilizing

shell, malachite green SERS signals from approximately

1–2 cm depth in the tissue could be detected using 785 nm

excitation light.11 The PEG-ylated SERS nanoparticles were

brighter than NIR emitting semiconductor quantum dots.

They were conjugated with tumor marker-specific antibody

fragments and subsequently detected in a tumor model.11

Whole body imaging of SERS labels in mice was also reported

for commercial, glass-coated SERS labels,12 and their systemic

toxicity was evaluated.65 Later on, the same group reported

imaging of ten different of these SERS labels upon subcutaneous

injection in ten separate areas.66 Multiplex imaging of the

liver, where five different SERS labels had accumulated after

simultaneous intravenous injection.66 Employing resonant

excitation of the reporter dye will add to the SERS enhance-

ment provided by the metal nanoparticles, and is useful

for in vivo detection. For example, mixed-monolayer gold

nanorods, coated with PEG polymers and visible- and NIR-

absorbing dyes could be used for multiplexed detection at

low toxicity.15,67,68 In vivo detection of these nanostructures

is very useful, as they are also used for photothermal tumor

therapy.67,68

Caenorhabditis elegans (C. elegans), a small, transparent,

well-characterized nematode49 (Fig. 2A), is a very convenient

complex system for spectroscopic studies, including whole

body imaging by infrared spectroscopy69 and stimulated

Raman scattering microscopies.70–72 C. elegans stores lipids

in granules in the intestine. The dye Nile red interacts with

such lipid granules through lipophilic or hydrophobic inter-

actions. When attached to silver nanoparticles, the nano-

particles are also targeted to the lipid granules, thereby

displaying the lipid droplets based on the SERS spectrum of

Nile red.50 Images can be generated in a confocal microscope,

setting the detected spectral region to the spectral range

covered by the SERS spectrum of Nile red (Fig. 2B).50 In

their study comprising fifty C. elegans samples, Charan and

co-workers showed that the silver nanoparticles lead to a

SERS signal from the lipids in their environment. This signal

can be imaged in a similar way as that of Nile red, by selecting

the spectral range of the CH stretching modes as an integral

signal for imaging (Fig. 2C). This is one of the very few

examples, where intrinsic SERS signatures from the tissue or

cell have been used for whole body Raman imaging. Small

animals with well-resolved microstructure can provide better

insights into the possible routing of nanomaterials, also the

distribution in different cell types. As another example, Wang

et al. described the distribution of gold nanoparticles that were

labeled with para-mercaptobenzoic acid (pMBA) in zebrafish

embryos using the SERS signal that is collected from the

different regions of the embryos and in different stages of

their development.73 The nanoparticles were micro-injected

into the one-cell stage. When a mixture of two kinds of gold

nanoparticles, one bearing pMBA, the other mercaptopyridine

were introduced into the embryo, the distribution was very

similar, as evidenced by the constant intensities of their

contributions in the spectrum.73

In order to follow the paths of nanomaterials by in vivo

Raman spectroscopy, small non-vertebrate model organisms

such as C. elegans provide a number of advantages over

mammals, since the penetration depth of the excitation and

scattering light will always include the whole animal. Further-

more, the application of higher magnification in a Raman

microscope could enable the retrieval of some histological

information or even data on subcellular localization in the

same experiment.

For studies of larger animals, for example small rodents in

the laboratory, Raman imaging of the whole organism may

require several different types of experiments. While confocal

approaches can be applied for the characterization of the

skin74 and subcutaneous regions11,66 spatially offset Raman

scattering (SORS) is a very promising technique for probing

deeper layers of tissue in the body.1,75,76 This has been

illustrated also in many studies on bone by Morris and

Matousek,77,78 including tomographic applications.79 Other

strong Raman scatterers, such as accumulations of SERS

active nanoparticles with characteristic spectral signatures

in deeper tissue layers, can be detected by SORS. First

proof of this was provided by Stone and co-workers,

who recovered Raman signals from SERRS tagged nano-

particles non-invasively from chunks of tissue,80 later also

from samples on the order of 45–50 mm thick by surface

enhanced spatially offset Raman spectroscopy (SESORS).81

Signals from four different types of commercial SERS tags

could also be imaged based on SESORS data.81 Van Duyne

Fig. 2 Whole body Raman imaging in the nematode Caenorhabditis

elegans. (A) Micrograph of C. elegans (reprinted with permission from

Jorgensen and Mango49). (B, C) Images of the lipid granules using

lipid targeting Raman probes consisting of silver nanoparticles and

Nile red using the signal of lipids (B), and of Nile red (C) (reprinted

with permission from Charan et al.50).

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5784 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

and co-workers combined the high detection sensitivity of

SERS with the in vivo potential of SORS for quantitative

transcutaneous glucose measurements in living rats.82 The

experiments were conducted with an implanted solid SERS

sensor without additional reporter molecules, out-of-resonance,

and they illustrate that SESORS imaging could also be used to

describe the native, intrinsic chemical environment of individual

nanoparticles or accumulations of nanoparticles in live animals.

Thereby, a SESORS approach could not only help to study the

distribution of nanomaterials in living animals, but also in

the description of the biochemical environment of the materials

in the organism.

As illustrated by the distribution of SERS active nano-

particles in the zebrafish embryos,73 developing systems,

organisms or individual differentiating cells provide particular

challenges for nanotoxicology research, as they display high

phenotypical, morphological and ultrastructural dynamics

that can result in physical re-distribution of nanostructures,

and changes in the cytotoxicity with developmental stage.

Accumulation in specific organs of the body implies the

interaction of a nanomaterial with individual cells. The work

cited so far did not discuss the interaction at the cellular level,

but the main portion of this article, to follow here, will focus

in detail on the interaction of the nanomaterials with the

cellular ultrastructure.

3 Nanomaterials inside cells

3.1 Endocytosis and endosomes

To study the reaction of cells to nanoparticle exposure and

how particles are internalized, the investigation of the uptake

mechanism is the first crucial step. In fact, the uptake of a

material will determine its fate in the cell, as it influences the

position in the cellular architecture, the routing within the

cellular ultrastructure, and the potential processing pathways.

In some, although few, Raman experiments, nanoparticle

injection was chosen to administer metal nanoparticles into

cells for SERS experiments, to avoid interaction of the nano-

particles with components from cell culture medium and to

achieve direct targeting of the cytosol,19 or to reach one

specific cell.73 Others have used electroporation for the direct

uptake of nanoparticles.84–86

In contrast, in the natural situation, nanomaterials enter

eukaryotic cells of animal origin using a variety of ways.87–90

The physicochemical properties of the nanomaterial, e.g. size,

shape, surface charge and modification, determine their

uptake efficiency and localization.91–94 Some carbon nano-

tubes were observed to directly penetrate the membrane of

cells, similar to needles,56,95 and also small PEG-functionalized

gold nanoparticles were observed to roam free in the cytosol,

suggesting their direct uptake.89 An important uptake

mechanism for nanomaterial is endocytosis. Endocytosis is

sensitively regulated, and serves both the uptake of nutrients

as well as signaling in the cell. By endocytosis, the cell can take

up macromolecules, liquids and pathogens, and also most types

of artificial nanoparticles.96,97 When the material approaches

the surface of the cell, the cellular membrane invaginates,

and forms a membrane vesicle inside the cell, the endosome.

The transmission electron micrograph in Fig. 3A illustrates the

invagination of the cellular membrane around a gold nano-

particle in an epithelial cell. Inside the cell, the endosomes,

including their cargo, are transported along a network of

actin filaments and microtubules. Endosome formation varies,

depending on the type of the cargo and on its size (Fig. 3B):

after interacting with surface receptors or specific regions in

the membrane of phagocytic cells, large, micron-sized particles

are taken up by phagocytosis, liquids by macropinocytosis.

The endocytic process of nanoparticles o300 nm is usually

mediated by proteins in the cell membrane, the most impor-

tant mediators being clathrin and caveolin.98,99 The nano-

particles interact with receptor molecules in the membrane, or

membrane rafts displaying a specific lipid composition. A

recent critical review discusses these and other mechanisms

associated with endocytosis and the processing of endosomes

in detail.96 Inside the cell, most of the nanomaterial resides in

endolysosomal structures. In Fig. 4, transmission electron

micrographs of late endosomal and/or lysosomal vesicles are

shown, containing an accumulation of gold, silver, and silica

nanoparticles, as well as DNA-functionalized SWCNTs. The

TEM micrograph displayed in Fig. 4F shows SWCNTs in

membrane-enclosed vesicles, which suggests an endocytotic

transport mechanism for these particular SWCNTs.9 In order

to reach other cellular organelles, such as the nucleus, the

endocytosed nanoparticles would need to escape from the

endosomal vesicles. Strategies for attaining endosomal escape,

as they are pursued, e.g., by viruses and bacteria have been

summarized in ref. 96.

Fig. 3 Uptake mechanisms of nanoparticles. (A) Endocytosis of a

gold nanoparticle by an IRPT epithelial cell (adapted from Kneipp

et al.24). (B) Schematic: depending on the cell type and material

properties, the particle uptake occurs via phagocytosis, pinocytosis

or receptor-mediated endocytosis.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5785

In the following sections we will illustrate how the process of

endocytosis and its effects on the cells can be studied by

Raman scattering. Specifically, the existence of endosomes

inside the cell has been evidenced by the following kinds of

experimental results:

(i) The accumulation and re-distribution of Raman spectro-

scopic signals that can be attributed to the nanomaterial,

e.g., ref. 9, 100, and 101.

(ii) Changes in surface-enhanced Raman scattering intensities

that indicate changes in the plasmonic properties of gold and

silver nanoparticles related to processing and aggregation in

aging endosomal vesicles.4,24

(iii) The ability to monitor endosomal transport of metal

nanoparticles over time.102,103

(iv) Raman-based measurements of pH in the environment

of gold nanoparticles, indicating pH values that are typical of

different endosomal stages and monitoring of these changes

over time.104,105

(v) Evidence of a biochemical composition that is charac-

teristic of processes associated with phagocytosis based on

resonance Raman data e.g., ref. 18.

(vi) Evidence of a biochemical composition, and molecular

processing that is characteristic of endosomes and lysosomes

based on normal Raman scattering and SERS e.g., ref. 24

and 106.

While items (i)–(iii) focus on spectral evidence that are

a direct consequence of the quality of the nanomaterials,

the latter three (iv)–(vi) comprise an interrogation of the

biological component, using the compositional and structural

information that is contained in the Raman spectra from

the cells.

3.2 Observing nanomaterials inside cells with Raman

spectroscopy

Many nanomaterials have specific optical properties that can

result in a higher Raman cross section, often due to resonant

Raman (RR) scattering. Low-dimensional carbon nanostructures,

in particular single-walled carbon nanotubes (SWCNTs) (Fig. 5A

and B), but also multi-walled CNTs, and single-walled carbon

nanohorns (SWCNHs) (Fig. 5C) can be observed in the cells

based on their strong resonant Raman spectra, the most

prominent example being SWCNTs.9,58,101,107 Other examples

of nanomaterials that can be detected by RR or normal Raman

scattering are drug or dye labeled, or unlabeled nanoparticles of

polymers (Fig. 5D–F), gold, silver, or silica.13,108–110 In the case

of silver or gold nanostructures, the nanoparticles provide high

local fields in their very close vicinity, and surface-enhanced

Raman scattering (SERS) leads to high Raman signals.16,111

The high intensity of spectral contributions from the molecular

or nanomaterial species in the RR or SERS case enables

probing of the nanoparticles’ properties inside the cells at low

concentration, and selective imaging of the nanomaterial.

Several studies report on the uptake and distribution of

SWCNTs in cells, evidenced by the strong and characteristic

RR spectrum.Heller and co-workers imaged the spectral signature

of SWCNTs encapsulated by an oligonucleotide and mapped

their localization in 3T3 fibroblast cells (Fig. 5A and B).9

Fig. 4 Nanoparticles in endosomes. TEM images of ultramicrotome slices of eukaryotic cells show metal nanoparticles (A–C), silica

nanoparticles (D, E) and SWCNTs (F) with respect to the cellular ultrastructure. Mouse fibroblast cells were incubated with gold nanoparticles

for 3 (A) or 24 hours (B). Silver nanoparticles (C) are enclosed in endosomes after 3 hours of incubation. (D, E) Silica nanoparticles were taken up

into 3T3 fibroblast cells after a 24 h-exposure (adapted with permission from Drescher et al.83). (F) TEM images of vesicle-bound DNA–SWCNTs

in murine myoblast stem cells (scale bar represents 1 micron; reprinted with permission from Heller et al.9).

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5786 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

TEM evidenced accumulation of the SWCNTs in perinuclear

lysosomes (Fig. 4F).9 A similar nanomaterial, single-walled

carbon nanohorns (SWCNH), was studied by Tetard and

colleagues.101 SWCNH provide strong and distinct Raman

spectra that can be clearly detected when the structures reside

in macrophage cells (Fig. 5C).

Also functionalization dependence of SWCNT uptake was

studied by Raman microspectroscopy. In particular, localiza-

tion of SWCNT carrying surface modifications with PEG

chains of different lengths was imaged.112 PEG-functionalized

SWCNTs were also studied by Raman microscopy in neuronal

PC12 cells.61

The group of examples for intracellular nanoparticulate

labels with distinct, strong Raman signatures comprises many

different types of SERS hybrid labels. They consist of different

gold or silver nanostructures, such as spherical nanoparticles,

nanorods, nanoshells and others, that are combined with so-called

reporter molecules, which provide a strong characteristic Raman

signature to the nanomaterial.

When SERS labels are used to highlight cellular structures,

analogously to dyes or quantum dots in fluorescence labeling,

they can be followed and imaged inside a cell based on the

SERS spectrum of their reporter molecule. Dye reporters

include indocyanine green,113 crystal violet,114,115 methylene

blue,109,114 rhodamine 6G,116 eosin,109 Giemsa109 and others.

Also ‘colorless’ molecules have been used as Raman reporters.

Often, they are attached to gold or silver nanoparticles by a thiol

group. Popular candidates are mercaptopyridine, aminothio-

phenol, para-mercaptobenzoic acid, or thiocresol.104,115,117–125

Since these molecules will change the surface charge of nano-

particles, they can in principle result in different uptake behavior

of the cells.119,126 In all these experiments, the distribution of

reporter SERS signals can provide an indication for the inter-

action of the nanostructure with the cell. For example, as

illustrated in Fig. 5H–J mixing of different SERS nanoprobe

populations carrying different reporters upon subsequent incuba-

tion with cells can be imaged using multivariate analyses.120

The uptake, accumulation and re-distribution of SERS labels

in cells can be monitored over the course of many hours and

days. In Fig. 6, the results of an experiment are shown where the

time-dependent uptake of gold nanoparticles and their intra-

cellular distribution were studied using SERS nanoprobes

functionalized with para-aminothiophenol (pABT). Here,

mouse fibroblast cells were incubated with citrate-stabilized

gold nanoparticles coated with pABT (100 nM) for 30 minutes.

After particle incubation, cells were washed thoroughly with

PBS buffer and covered with fresh cell culture medium. In

principle all cell samples were exposed to the same amount

of nanoparticles for the same time. SERS nanoprobes were

internalized into 3T3 fibroblast cells by passive uptake via

fluid-phase. The Raman measurements were conducted after

different time-points ranging from 30 minutes to 5 days

(schematic in Fig. 6A).

Raman intensity maps and SERS spectra of single live cells

incubated with pABT gold nanoparticles are depicted in Fig. 6B.

To construct these images, SERS spectra were acquired by raster

scanning over an individual cell in steps of 2 mm. The Raman

intensity maps in Fig. 6B are representative for each time-point

and illustrate the local distribution of the pABT gold nanoprobes

over the entire cell. To visualize the intracellular SERS nano-

probes, the relative signal intensity of the C–S stretching

vibration of pABT at 1085 cm�1 is plotted. Already after

30 minutes of incubation, pABT is detected inside the fibro-

blast cells in the close proximity to the plasma membrane.

With increasing incubation time (3 hours to 2 days), particles

Fig. 5 Confirming the presence of nanomaterials inside cells using

Raman scattering. (A) Combined Raman and fluorescence spectra of

DNA-SWCNTs in live 3T3 fibroblast cells after 8 days in culture. The

corresponding Raman intensity map of the radial breathing modes of

nanotubes in 3T3 cells is shown in (B) (scale bar: 20 micron; reprinted

with permission from Heller et al.9). (C) Raman spectra of single-

walled carbon nanohorns (SWCNHs) (black spectrum) and SWCNHs

within a macrophage (red spectrum, L1) 7 days after exposure

(reprinted with permission from Tetard et al.101). (D–F) High-resolution

Raman imaging of a phagocytosed latex bead in a neutrophil (Copyright

2005 National Academy of Sciences, USA). Raman intensity maps (6.7�6.7 mm2) of the 1000 cm�1 band of polystyrene (D) and a corresponding

cluster analysis image (E) are shown. The average Raman spectra are

extracted from the blue (spectrum 1), magenta (spectrum 2) and yellow

(spectrum 3) clusters displayed in (F). Spectrum 1 shows a strong signal

of polystyrene (reprinted with permission from van Manen et al.108).

(G–J) Duplex imaging using two different SERS hybrid labels in a 3T3

fibroblast cell (reprinted with permission from Matschulat et al.120.

Copyright 2010, American Chemical Society). (G) Photomicrograph of

the cell (scale bar: 4 micron), (H) chemical image, based on one intensity

from each of the labels (I, J) images obtained using hierarchical cluster

analysis and K-means clustering, respectively, with the spectral informa-

tion in the region 300–1700 cm�1, based on three spectral classes.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5787

are evenly distributed in the cytosol and accumulate in the

perinuclear region in the course of endo-lysosomal processing.

The Raman intensity map acquired after 2 days of incubation

do not show signals in the region of the cell nucleus. After

5 days of incubation and several cell divisions the intracellular

signal intensity of pABT and the amount of detected Raman

spectra per cell decrease significantly since the primary vesicles

are distributed over more cells. During cell division, the

cytoplasm is divided between the two daughter cells. This dilution

of SERS nanoprobe-containing endosomes is a trivial effect on cell

lines, and has been observed in other studies as well.126 However,

observing particle distribution between differentiating cells in

developing tissues may help to understand better the distribution

of cytoplasmic and nanomaterial material in developing

organisms,50,73 as was already discussed in the context of

whole body Raman imaging in Section 2.3.

Selected SERS spectra of different stages of the endo-lysosomal

pathway are illustrated in Fig. 6C. Independent of the incubation

time, pABT is identified in each spectrum. The spectra of the

shorter incubation times are dominated by characteristic Raman

bands of pABT. After 1, 2 and 5 days of incubation, cell-specific

Raman bands are detected in addition to the pABT Raman

signature. This is attributed to the replacement of particle-bound

pABT by cellular molecules with increasing incubation times. Since

the Raman spectra show the same pABT signature after 30 minutes

and 5 days, we can assume that pABT is stable during cellular

processing. Furthermore the signal intensity is increased from

30 minutes to 48 hours of incubation, which is correlated with a

higher SERS enhancement. This can be assigned to changes of the

particle aggregation with increasing incubation times due to multi-

vesicular fusion (see also TEM images of gold nanoparticles in

Fig. 4). The SERS nanoprobes used in these experiments induce

no cytotoxic effects as evidenced by XTT assay.120

We would like to point out the difference of intracellular

application of SERS labels to the many experiments that are

done with SERS labels in immuno-cytochemistry, mostly to

detect cell surface receptors in individual cells or antigens that

become exposed after fixation of a tissue.127–134 Similarly, we

have omitted to include a discussion here on the possibilities of

using SWCNTs for labeling, as shown in e.g., ref. 135. We

have not included these immunocytochemistry studies in our

discussion here, as they involve the targeted interaction of the

nanomaterial with the surface of cells for diagnostic purpose,

determined by an antibody or peptide, and largely independent

of the properties of the nanomaterial.

3.3 Modification of nanomaterials inside the cells

The biological system can exert great influence on the physico-

chemical properties of a nanomaterial, e.g., by chemical reactions

taking place at the surface of nanoparticles, by adsorption of

molecular species, resulting in changes of surface charge,

solubility, and aggregation behavior. Changes in nanoparticle

surface modification can be a result of processing of the nano-

material by the cell. Considering the function of endolysosomes

as the cell’s waste disposal and the abundance of proteases and

other hydrolyzing enzymes taking care of the degradation of

biomacromolecules, it seems obvious that nanomaterials will

be modified in these cellular compartments as well. Raman

experiments employing SERS provide evidence of molecular

surface modification of gold and silver nanoparticles. For

example, Oyelere et al. introduced peptide-modified nanorods

into cells, where they monitored the intensity of the C–N

stretch band of a thioalkyl-triazole linker for a targeting

peptide at 760 cm�1.106 From a decrease in intensity

they conclude that the conformation of the linker must

Fig. 6 Time-dependent study of nanoparticle uptake and their intracellular distribution by the use of SERS nanoprobes. (A) Experimental

design. Mouse fibroblast cells were exposed to gold nanoparticles coated with para-aminothiophenol (pABT, 100 nM) for 30 minutes. Raman

measurements were conducted at different stages of incubation. (B) Corresponding Raman chemical maps taken at a step size of 2 micron visualize

the distribution of nanoparticles inside the cells (using the intensity of the n(C–S) vibration of pABT at 1085 cm�1). (C) SERS spectra of

intracellular pABT gold nanoparticles after different time-points. An excitation wavelength of 785 nm was applied in all experiments with a laser

intensity on the sample of B8 � 104 W cm�2 and an acquisition time of 1 s.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5788 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

change, when the nanoparticles with the attached peptide

enter the cells.106

Similarly, the molecules that are used as reporters in SERS

labels may be modified or displaced from the nanoparticle

surface. Sirimuthu and co-workers reported the degradation

of the signal from reporter molecules on the surface of silver

nanoparticles, depending on whether the reporter molecules

were attached by chemisorption to the silver by a mercapto

function, as with 4-mercaptobenzoic acid, or by physisorption,

as in the case of the dye Rhodamine 6G.121 They observed the

signal of Rhodamine 6G physisorbed to silver nanoaggregates

to disappear after 8 hours in the cell culture medium, and

after B12 hours in Chinese hamster ovary and MG-63 cells.

It is remarkable that the SERS signature from the dye was

not replaced by SERS signals from the cellular molecules

could be detected instead in these experiments but disappeared

completely,121 contradictory to work with labeled and unlabeled

silver nanoparticles in cells, where signals from cellular molecules

have been reported, as will be discussed in detail below.4,119

In phagocytes, which can take up particles in the micron-

size range, drastic changes can be observed in the Raman

spectra of polymer materials that are being ingested. Using

RAW 264.7 macrophage cells, van Apeldoorn et al. could show

that large micron-sized particles of the polymer poly(lactic-co-

glycolic acid) (PLGA) were degraded in a heterogeneous way

B1–2 weeks after internalization into phagosomes, with a rapid

degradation in the center as compared to the outer parts.100

PLGA has a Raman spectrum with intense bands at 1768 cm�1

characteristic of ester bonds, and several bands that belong to

glycolic acid at 845, 890, 1274, and 1424 cm�1. Based on the

PLGA Raman spectra van Apeldoorn and co-workers showed

that ester hydrolysis occurs throughout the phagocytosed

particles, with a selective loss of glycolic acid units. They also

showed that molecules from the cell, such as proteins and

lipids, may travel into voids that are brought about by the

degradation process inside the spheres.100 Degradation of

microspheres from other materials inside RAW 264.7 cells

was studied by the same group.136 This work includes the

quantification of dextran in phagocytosed cross-linked

dextran hydrogel microparticles using an intensity ratio from

their Raman spectrum, and led to the result that dextran

particles degrade by swelling. In contrast, poly(ethylene glycol)-

terephthalate–poly(butylene terephthalate) (PEGT–PBT) block

copolymers did not show changes in their Raman spectra when

ingested by the macrophage cells.136

Degradation of polymer particles can follow drastically

different patterns depending on their size, on the cell type

and uptake. In contrast to the micron-sized PLGA spheres

studied in macrophages by van Apeldoorn et al.,100 PLGA

nanospheres that were endocytosed from liquid culture medium

by non-phagocytosing HeLa cells were found in the perinuclear

region B2 hours after incubation, suggesting their interaction

with the endoplasmic reticulum and Golgi apparatus.13 When

the incubation with the PLGA nanospheres was followed by

chase incubation with regular culture medium, degradation of

the nanoparticles was found after 3 hours, as indicated by lipid

and phospholipid spectral contributions, in particular by the

frequency change in the carbonyl stretching vibration that

shifted from the frequency characteristic of the PLGA ester

linkage around 1765 cm�1 to lower frequency typical for lipids

and phospholipids B1740 cm�1.13 In a similar fashion, the

Raman signals from poly-caprolactone (PCL) nanoparticles were

reported to decrease concomitantly with an increase in spectral

features that could be attributed to membrane structures possibly

belonging to late endosomes associated with the Golgi.13

Depending on the nanomaterial and the basis of its char-

acteristic Raman spectrum, the influence of the biosystem on

the Raman spectra of the nanomaterial can vary greatly.

Changes due to the interaction of the nanoparticles with

components in the cellular environment can be more subtle

when the material is not chemically degraded. In some materials,

optical properties may change drastically with their aggregation

due to accumulation in endosomal structures. This will be

discussed in the following using the examples of SWCNTs and

of plasmonic nanoparticles.

The Raman spectrum of SWCNTs is determined by the strong

coupling between electrons and phonons of the nanotube,

occurring if the incident or scattered photons are in resonance

with an electronic transition between van Hove singularities in

the valence and conduction bands. The resonant Raman spectra

are very sensitive to the nanotube diameter and chirality,58 and

Raman scattering can be used to assess these properties in single,

isolated SWCNTs.107 In some types of SWCNT bundles, the

typical band structures may be altered through close nanotube–

nanotube parallel contacts.137 In the resonant Raman experiment,

the modified resonance conditions change the observed specific

radial breathing modes (RBM) of the nanotubes138,139 in the

low-frequency region of the SWCNT spectrum.58

The possibility to characterize isolated SWCNTs enables

their observation inside biological cells and their interaction

with other SWCNTs. An example illustrating this is the

application of chemical stains to cells that incorporated

SWCNTs. Heller and co-workers found that when 3T3 mouse

fibroblast cells that had endocytosed SWCNTs and were

subsequently stained with hematoxylin and eosin, the RBMs

of the SWCNTs displayed an intensity increase in one RBM at

267 cm�1, indicative of a shift in the interband transitions.9 At

the same time, the fluorescence decreased, leading the authors

to conclude that the SWCNTs must be in direct contact.9,138

Since the characteristic RBM contribution at 267 cm�1 had

been absent in unstained cells, the staining procedure that

involves several chemicals was held responsible for causing

SWCNT–SWCNT contact. Even though this was induced by

externally applied stains after endocytosis of the SWCNTs, the

example illustrates that chemical modification of SWCNTs in

harsh lysosomal environments could in principle be detected in

the Raman spectrum. When PEG-functionalized SWCNTs

were studied ex vivo in mouse tissues in liver macrophages,

equal concentrations of oxidized SWCNTs gave lower signals

than unoxidized SWCNTs due to covalent damage to the

carbon atom networks in the oxidized sample.60

In the case of gold and silver nanoparticles, not their own

Raman spectrum is usually observed, but the SERS spectrum

of molecules in their close proximity. High SERS signals are mainly

caused by high local optical fields around the nanostructures.140–145

The dependency on the local fields implies that the SERS

enhancement strongly depends on the morphology (e.g., the

size, shape, or aggregation) of the nanoparticles. Therefore, the

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5789

spectral information that can be obtained of the biological

component is coupled very strongly to the physicochemical

properties of the nanoparticles. In a SERS experiment, this

strong connection expresses in the following characteristics:

(i) the observed SERS intensities vary with the enhancement

factor, which depends critically on the ability of the nanoparticles

to form aggregates,146,147 (ii) the enhancement depends on the

type of aggregates, particularly on the gap size between nano-

particles and their geometries,148–151 and (iii) the average intensity

will depend on the number of nanoparticles or nanoaggregates

in the probed volume (compare also Fig. 6B). Therefore, even

if the Raman spectrum may not provide direct spectral

evidence of the material itself, the SERS data can be used to

gain information on the behavior of nanoparticles, such as

aggregation and re-distribution in a cell.

To illustrate the first two of the issues itemized above, the

results of an experiment are shown, where gold nanoprobes

were applied in the epithelial cell line IRPT.24 The nano-

particles were added to cell culture medium and removed after

30 minutes. Then, in a series of samples taken at different

time points, the SERS signal generated in the vicinity of the

nanoprobes was observed (Fig. 7A–C). We found the signal

strength of the SERS to vary significantly for different time

points after the endocytosis of the nanoparticles had taken

place. This gives information on the SERS enhancement

and thereby indicates that the morphology of the gold nano-

structures inside the endosomal vesicle must have changed.

Specifically, aggregates of nanoparticles must have formed24

(compare item (i)). In accord with theoretical predictions we

have shown recently experimentally that the enhancement of

individual gold nanospheres is very low around 102, but that

the formation of aggregates can change the enhancement factor

by many orders of magnitude.147

Item (ii), the signal strength as a function of nanoaggregate

morphology and aggregation was investigated by a TEM

study carried out in parallel samples.24 In our experiments,

the spectra obtained after 120 min (Fig. 7B) exhibited the

strongest signals. This improvement in signal strength correlates

with the formation of small gold aggregates, at that time mostly

dimers and trimers.24 TEM images indicated that later in the

time course after 180 minutes, inter-particle distances in the

gold nanoparticle accumulations were greater than in the

dimers and trimers observed after 120 min (see the arrow in

Fig. 7D). The higher interparticle distances, which are probably

due to multivesicular inclusions, may lead to a decrease in the

overall SERS intensity again (Fig. 7C).

Even though many nanomaterials do not show plasmonic

properties and cannot be used as SERS probes, understanding

better the interaction of silver and gold nanostructures with

cells will provide significant progress, considering that silver

nanoparticles have become ubiquitous in our everyday life yet

can display high cytotoxicity.152–155 Furthermore, the plasmonic

nanoparticles might serve as model systems to a certain extent, as

some general principles about the interaction of nanoparticles

with cells may hold for other materials as well.

4 Spectral evidence of the biological component

In many applications, Raman scattering has been used to observe

nanoparticle-bio-interactions based on the Raman spectroscopic

information from the nanoparticles. This is probably the case

because questions posed by biotechnology, imaging, and the

need for more efficient labels have served as a major driving

force for research in the area of nano-bio-Raman. One should

be aware that in some experiments, changes in the properties of

the material are difficult to detect based on the Raman spectrum

of the material itself. This can be due to insufficient sensitivity to

minute amounts of nanoparticles in the biomatrix, due to the

different conditions under which Raman experiments on cells

and tissues take place compared to experiments on some

inorganic materials, or a broader, indistinct Raman spectrum

that would become less obvious in the context of sharp bands

from the biological molecules in its surroundings, e.g. in the case of

amorphous silica nanoparticles.156 In such samples, observing

changes in the biochemical composition, in the molecular processes

and interactions within the cellular architecture becomes of vital

importance for delineating the influence of the nanomaterials.

In comparison to approaches in molecular biology that

usually rely on a specific component of the cellular systems,

e.g., protein expression patterns, generation of ROS, changes

Fig. 7 (A–C) Typical SERS spectra from cells of the epithelial cell line IRPT after incubation with gold nanoparticles, excited witho3 � 105 W cm�2

at 785 nm, collection time 1 s. The incubation time, including a 30 min nanoparticle pulse, is indicated in each panel. (D) Transmission electron

micrograph of IRPT cell showing gold nanoparticles in endosomal structures. Arrows point at large distances between the nanoparticles, probably

brought about by multivesicular inclusions (scale bar: 250 nm). (Reprinted with permission from Kneipp et al.,24 Copyright 2006, American

Chemical Society).

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5790 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

in metabolic activity, or proliferation of the cells, the Raman

spectroscopic fingerprint provides a combination of all the

‘phenomic’ information from all molecules that are contained

in a probed sample volume.

4.1 Chemical composition of phagocytic cells in the presence of

nanomaterials

Phagocytic cells are an important cell type, considering that

nanomaterial which enters an organism is likely to encounter

cells of the immune system, which are specialized in ingesting

foreign materials. Different uptake mechanisms are responsible

here than in the receptor-mediated endocytosis observed for the

smaller nanoparticles, and large entities can be taken up,

including large particles and agglomerates (Fig. 3). Phagocytes

such as macrophages, granulocytes and others engulf ingested

material, usually pathogenic particles inside their phagosomes,

and then destroy the phagocytosed particles. An enzyme

complex, NADPH oxidase is needed to produce oxidants for

the degradation of the foreign material. When the phagocyte is

active, the enzyme complex assembles on the phagosomal

membrane to produce reactive oxygen species, and is associated

with a cytochrome unit, flavocytochrome b558.

Puppels, Otto, Sijtsema, van Manen and co-workers

have pursued a great number of studies on phagocytic cells,

comprising Raman spectra of heme containing peroxidases,

cytochrome b558 in different types of granulocytes,18,157,158

lipid and protein composition,108,159 and foam cell formation

of macrophages.136 In one of the first papers on this issue,

Puppels et al. stated, ‘‘monitoring the phagocytotic process

and digestion of foreign objects {would be an} . . .exciting

prospect’’.157 Meanwhile, in fact several new observations

were made in the process of phagocytosis using non-resonant

and resonant Raman microspectroscopy, and many aspects

have been investigated.18,108,157–159 More recently, Chang

and co-workers reported the quantitative Raman microscopic

observation of the phagocytic degradation of ingested yeast

cells, along with a temporal characterization of phagocytic

NADPH oxidase.160

In Fig. 5D and E, we had shown the work of van Manen

and co-workers, who had observed Raman spectra of latex

microparticles in neutrophilic granulocytes.108 Raman spectra

measured in the proximity of these phagosomes revealed also

the presence of lipid bodies, which contained high contributions

from arachidonate.108 Differences between the lipid bodies

of cells deficient in a NADPH oxidase subunit and wild type

cells were found, which led to the conclusion that this flavo-

hemoprotein mediates the assembly of the lipid bodies around

the phagosomes.

Macrophages, upon ingestion of micromaterial, can convert

into foam cells, that is, cells that are filled with lipid droplets.

Their high content in unsaturated lipids and cholesterol

and cholesterol esters was also shown by non-resonant

Raman imaging.136 The relatively large body of Raman

literature on the biochemistry of phagocytosis illustrates

that in addition to spectroscopy and imaging technologies,

systematic work with suitable, consistent in vitro models

may pay off, when a specific aspect of nano-bio-interaction

should be clarified.

Differences in the SERS spectra obtained from the mole-

cules in the enhanced local fields of nanostructures over

time have shown that SERS approaches are feasible for the

characterization of changing cellular environments and useful

for intracellular applications.16,24,102 Using SERS spectra

generated due to the presence of endocytosed gold nano-

particles, the endosomal composition in phagocytic cells can

be studied.

When gold nanoparticles were added to the culture medium

of the macrophage cell line J774, and spectra were measured

at different time points after incubation (60 min, 120 min,

180 min), a great variety of SERS spectral patterns was found,

indicative of many different molecular species present in the

endosomes of the macrophages, that changed with incubation

time.24 Nevertheless, one specific spectral signature appeared

at all time points, with exception of the very late, lysosomal

stage (22 hours incubation). The spectrum is displayed

in Fig. 8, together with a bright field image of J774 cells

containing large endolysosomal accumulations of gold nano-

particles, displaying as black spots. Almost all bands in the

spectrum are characteristic of adenosine phosphate and show

contributions from all constituents of this nucleotide (adenine,

ribose, and phosphate161). A comparison with SERS and

normal Raman data of pure adenosine monophosphate

(AMP) and adenosine triphosphate (ATP) from the literature

suggested a prevailing contribution from AMP, since triphos-

phate and diphosphate markers are lacking. Interestingly, the

acidification and pH regulation of endosomes are achieved by

an ATP-dependent proton pump and a Na–K-ATPase.162,163

Furthermore, the endosome acidification profile can be

modified by cyclic AMP.164 The outcome of this experiment

suggested that in fact molecular species involved in the

generation of a specific endosomal milieu can be detected

and observed by SERS.24

Fig. 8 SERS spectrum collected from a spot in a J774 macrophage

cell after incubation with gold nanoparticles, excitation wavelength

o3 � 105 W cm�2 at 785 nm, collection time 1 s. The spectrum

represents the SERS spectrum of AMP and/or ATP. Band assign-

ments based on ref. 161 are indicated. Abbreviations: cps, counts

per second; n, stretching mode; o, wagging mode; d, bending mode;

Rib, ribose; Pyr, pyrimidine; Im, imidazol (adapted with permission

from Kneipp et al.24). Inset: photomicrograph of J774 macrophage

cells after incubation with gold nanoparticles.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5791

4.2 Endosomal constitution in the presence of the

nanomaterials: insights from SERS

SERS is an ideal tool to study endosomal composition because

the dimensions of the endosomes fit the dimensions of nano-

particles and their aggregates. In such an experiment, noble

metal nanoparticles that show plasmonic properties have to

be employed, since the endocytosed material itself serves as

the SERS-active nanostructure. Acting as a nanosensor, it

transmits molecular information from its immediate vicinity,

whose dimensions are determined by the local optical field that

is generated due to excitation of the localized surface plasmons

in the metal. SERS can probe the endosomal environment very

locally. In accord with ultrastructural studies using electron

microscopy, confirming accumulation of most endocytosed

gold and silver nanoparticles in endosomes (see also

Fig. 4A–C and Section 3.1), most work on SERS inside cells

provides information on the composition of endosomal

structures.24,102,113,114,116,165–168

It should be noted that the SERS enhancement can differ

drastically between individual nanoparticles and nanoaggregates

that appear as a consequence of vesicular fusion inside the cells

(see also Section 3.3). In systems where individual nanoparticles

prevail, probing the endosomal environment with silver nano-

particles rather than gold nanoparticles could be beneficial: even

though the enhancement of silver nanoparticle aggregates and

gold nanoparticle aggregates is very similar (both can reach up to

1014), the enhancement of individual silver nanoparticles

(106–107) is much higher than that of gold nanospheres

(103–104) at their respective plasmon resonances.145 On the other

hand, silver nanoparticles may be toxic to the cells at high

concentration, and may thereby affect endosomal composition.

Therefore, care should be taken when selecting plasmonic

nanomaterial for probing of early endosomal stages.

SERS spectra from cellular molecules were reported in

several different cell types, using either only gold nano-

particles,24,102,165,167,169 or gold or silver nanoparticles carrying

reporter molecules in a concentration that permitted the detec-

tion of intrinsic SERS spectra from the cellular biochemistry in

addition to the reporter signature.113,114,116,120,168

An assignment table for SERS spectra from endosomes is a

smorgasbord of bands, representing all types of molecules

present in the vesicles (for examples please see the works

cited above).

The detection of typical adenosine monophosphate signatures

in endosomes of J774 macrophages discussed above (Fig. 8)

illustrates though that important molecular species may be

identified in the spectra. Sorting of the spectral information from

SERS experiments in complex molecular mixtures is one of the

main tasks that will need to be accomplished in the current efforts

to establish intracellular SERS as a tool in biophysics and

nanotoxicology. Using some sort of statistical analysis can help

to pre-sort spectral signatures.

Coming back to an example discussed above in another

context, Fig. 7A–C displays typical SERS spectra for three

different time points after a period of endocytotic uptake of

nanoparticles into IRPT epithelial cells from a cell culture.

Without attempting an interpretation, the differences between

the spectral signatures indicate that the molecular composition

of the endosomal vicinity of the nanoparticles changes over

time. To compare potential qualitative alterations, full data

sets of B500–800 spectra were analyzed per time point. The

number of bands in the spectra that display 90–100% of the

maximum signal level of each data set (time point) totals 2 to 8

bands at 30 min. A significant increase in the number of

spectral bands is found at t = 120 min. At this time point,

the portion of SERS spectra exhibiting 90–100% of the signal

level contains 13–19 characteristic bands, resulting from an

abundance of different spectral contributions that were not

observed for the other time points. From the comparison of the

120 min and the 180 min spectra (Fig. 7B and C, respectively),

we see that there occurs again a decrease in the number of

spectral features (3 to 6 bands in the SERS spectra of the

90–100% highest signal level). This or similar information

could be used for imaging and correlated with complementary

information, e.g., from electron microscopy.

Other possible approaches could be the decomposition of

signals by various means. In our very first work on SERS

hybrid probes, we were able to subtract characteristic spectra

of the reporter molecule indocyanine green from the hybrid

spectra, yielding spectra with the endosomal contributions.113

Recently, we separated contributions from two different reporter

molecules and those of different intrinsic cellular signatures using

principal components analysis (PCA) and used cluster methods

for imaging (see also Fig. 5I and J).120 This result indicates that

fast multivariate evaluation of whole sets of SERS spectra is

feasible, also for varying signal-to-noise ratio.120 Other examples

of multivariate decomposition of SERS data include the applica-

tion of singular value decomposition of SERS spectra from

mixtures of reaction products170 and use of cluster analysis on

intrinsic SERS signals of biological cells in extracts of pollen

samples.171 Multivariate methods, as shown for hybrid SERS

probes carrying reporter signatures and spectral contributions

from intrinsic molecules, will also enable more efficient imaging

of endosomal structures in the future.

The lipid composition in the environment of endocytosed

nanoparticles has been a subject of both SERS as well as

normal Raman studies. Considering that the lipids of the

cellular membrane are key components in the endocytosis,

and lipid deposits are associated with material uptake and

degradation in phagocytes (see Section 4.1). Raman spectro-

scopic evidence of lipids in the proximity of endocytosed

material shall be discussed here briefly as well. SERS spectra

of lipids were found for SERS hybrid probes consisting of gold

nanoparticles and crystal violet as reporter dye in some

positions of 3T3 fibroblast cells after an incubation of the

nanoprobes for two hours.114 Comparing these results with

earlier studies of the same types of nanoparticles that included

a TEM study, efficient interaction of dimers and trimers of

nanoparticles24 with the endosomal membranes must take

place. Interestingly, intracellular SERS can also be used

to detect the production of lipids shortly after the onset

of adipogenesis in endosomes of human adipose-derived

stem cells.169 In this context, also the ability to observe the

lipid SERS signals in the intestinal granules of C. elegans50

discussed earlier should be mentioned one more time (compare

Fig. 2C). Although not further explored in this study, apart

from imaging, molecular investigation of lipid bilayers could

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5792 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

provide more detailed insight into the composition of the lipid

granules.

The observation of lipids in the proximity of the endocytosed

nanomaterial that was made in SERS experiments using plasmonic

nanoparticles are corroborated by normal Raman experiments

with cells that endocytosed other nanomaterials: Chernenko

and co-workers observed the co-localization of poly-caprolactone

nanoparticles and vesicles of high lipid content in HeLa cells.13

PCA identified the local environment of polystyrene nano-

particles in A549 epithelial cells as rich in lipidic signatures

which were associated with localization of the nanoparticles in

the endoplasmic reticulum.172 In the work of Romero et al.

with oxidized carbon nanotubes, spectra of cellular lipid were

found together with those characteristic of the nanotubes in

HepG2 cells.173 In general, normal Raman microspectroscopy

is a powerful tool to study lipid composition and structure

in cells, as shown in several other examples e.g., ref. 174.

Tip-enhanced Raman scattering (TERS) before endosome

formation could provide another means for probing cellular

membrane biochemistry by plasmon-enhanced Raman scattering.

In this way membrane structure could be probed at positions that

may end up as endosomal membranes as a consequence of

interaction with nanomaterials. After proofs-of-principle that the

method can be used to study membrane composition175,176 and

that TERS spectra can be acquired in liquids,177 such experiments

have come within reach.

4.3 Information on endosomal pH from a Raman experiment

A parameter that varies drastically over time in the endosomes

containing nanoparticles is the pH value.178 In many cell types,

significant acidification compared to earlier endosomal stages takes

place in the lysosome. There, to ensure e.g., proper enzymatic

function, pH can be well below 5, and even lower (o4).179

The SERS signatures of particular reporter molecules as

part of SERS probes were found to be pH-dependent. SERS

spectra of 4-mercaptobenzoic acid (pMBA) on silver and gold

electrodes and the dependence of the spectra on the pH value

have been reported and discussed in terms of adsorption

geometry on the metal and on the state of dissociation of

the carboxyl group as a consequence of surrounding pH.180

The pH dependent SERS spectra of pMBA adsorbed on gold

nanoshells bound to a silicon substrate were used to create a

pH meter working over the range of 5.8 to 7.6 pH units.181

Using 4-mercaptopyridine as a reporter, the pH range detected

on the nanoshells can be extended from 3 to 7.182 Similar

experiments demonstrated that hollow gold nanospheres with

pMBA were responsive over a pH range of 3.5 to 9.183

SERS studies on silver nanoparticle clusters functionalized

with pMBA showed that the spectrum is sensitive to pH

changes in the surrounding solution in the range of 6–8.122

The authors concluded from a SERS spectrum measured from

these silver particles incorporated into Chinese hamster ovary

cells that the pH in the environment of the particles was below

6, an observation consistent with the particles being located

inside lysosomes.122 Studies with pMBA on gold nanospheres

demonstrated that pH in the endosomal system varied between

6.8 and 5.4 in endosomes of different ages.104 Meanwhile,

several experiments determining pH using reporter functionalized

nanostructures in cells have been reported, including pMBA

on gold nanoshells,184 pMBA fiber optic nanoprobes for intra-

cellular measurements,185 and also aminothiophenol on silver

nanoparticles186 and on gold nanorods.187

The application of a pMBA-based SERS nanosensor for pH

imaging of single live cells is illustrated in Fig. 9. In these

experiments the pH nanosensor consists of citrate-stabilized

gold nanoparticles covered with a monolayer of pMBA. The

SERS spectra in Fig. 9A show the characteristic pMBA

signature at different pH values ranging from pH 3.8 to 10.

The Raman bands at 849 cm�1 and 1424 cm�1, which correspond

to vibrations involving the carboxyl group, are sensitive to changes

in the pH value.180 The deprotonation of pMBA at higher pH

values leads to an increase of the signal intensity of the carboxyl

stretching vibrations (Fig. 9A). For calibration of the pMBA

nanosensor, the signal ratio of the (C–C)ring stretching vibration

at 1076 cm�1, which is not influenced by changes in the pH value,

and the COO� stretching vibration at 1424 cm�1 are used.

The mobile and biocompatible pH nanosensor allows us to

monitor the distribution of the pH value in single cells at

subendosomal resolution. In the results displayed in Fig. 9C,

3T3 mouse fibroblast cells were exposed to pMBA-coated gold

nanoparticles for 5 hours. In the Raman microspectroscopic

cell experiments, spectra were acquired by raster scanning over

a single live cell in steps of 2 mm. The SERS spectra measured

in the cellular environment of the pH nanosensor (Fig. 9B)

exhibit different pMBA signatures, which indicate different pH

values inside the cell.

The Raman chemical maps of individual fibroblast cells in

Fig. 9C visualize the intracellular distribution of the pH value.

Obviously, after a few hours of incubation, the incorporated

gold nanoparticles have accumulated inside the cell. Depending

on the localization of the pH nanosensor, the intraorganelle

pH value differs within the cells. Close to the cellular

membrane the pH value is higher than in the perinuclear

region. This is attributed to the maturation of the vesicles

containing the pH nanosensor. In the course of cellular

processing the pH value becomes more acidic from pH 7.4

in the endocytic vesicle formed during particle uptake to

pH 4.5–5 in the lysosomes. The latter often accumulate near

the cell nucleus. Since particles are continuously taken up

during the incubation time of 5 hours, early endosomes and

lysosomes coexist in the same cell. Ultrastructural studies of

this cell line (see Fig. 4) prove that at this stage, many particles

are enclosed in late endosomes and lysosomes near the cell

nucleus.

The distribution of sampled spots of different pH in the

cells, as displayed in Fig. 9C, can also be analyzed statistically,

and can be used to follow the dynamics of the pH change in a

population of endosomes.105

We also demonstrated the application of a pMBA SERS

sensor using two-photon excited Raman scattering (surface-

enhanced hyper Raman scattering, SEHRS).104 SEHRS spectra

of pMBA show the same bands as SERS spectra, but the

relative signal strengths are changed. These differences in

the SEHRS signature of pMBA are useful for creating a

pH sensor that allows measurements in a more extended range,

from pH 2 to pH 8.104 This could be of interest for measure-

ments in subcellular compartments of unknown or extreme pH,

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5793

e.g., very acidic lysosomes, and pathologies associated with

potentially defective acidification.188–190

4.4 Raman information from the nucleus

The nuclei of animal cells represent the largest sub-cellular

compartments and can be identified in bright field images of

unstained cells. Therefore, they can be studied easily using a

Raman microscope, and have been investigated frequently

since the early days of single cell Raman spectroscopy. Raman

spectra from intact nuclei can reveal information about the

in vivo secondary structure of the DNA and about its inter-

action with proteins.21,157

Since the DNA structure and association with proteins

change in the course of the cell cycle, the DNA and protein

spectral signatures from the nuclei have provided useful mar-

kers for the discrimination between different cell types and for

cancer cell detection in the development of Raman-based tumor

diagnostics, e.g., for the identification of neoplastic cells.76,191

The nucleus of a cell can be imaged either by mapping the

intensity of DNA-specific Raman bands, or by hyperspectral

imaging, the latter taking into account wider spectral regions or

the Raman spectral fingerprint as a whole.22,192,193 Imaging

studies using the resonant Raman signals of drugs have revealed

the specific interaction of externally applied molecules with the

nuclear ultrastructure. For example, teraphthal, a transition

metal complex that could serve as a component in a cytostatic

drug, was monitored in living A549 lung epithelial cells.194 RR

imaging revealed that the teraphthal binds very weakly to

DNA and RNA, but it readily forms complexes with the

histone proteins in the nucleus.194 In another, more recent

study, a CO-releasing metal–carbonyl complex was imaged after

its penetration of an HT29 colon cancer cell, and found to

localize mainly in the nuclear membrane and the nucleolus.195

Cytotoxicity, potentially induced when nanoparticles enter

a cell, can induce severe molecular changes in the nucleus. In

the example of silver nanoparticles, concentration dependent

cytotoxicity is observed,152,155 expressing as mitochondrial

and DNA damage or early apoptosis. Resonance Raman

data can report on the mitochondrial integrity. Recently,

Okada et al. observed dynamic changes in the cytochrome

c distribution inside cells after inducing apoptosis.23 Their

data using resonant Raman excitation of cytochrome c suggest

that the distribution of this protein changes because the

molecule is released when mitochondrial integrity is lost in

the apoptotic process. The work also showed that the redox

state of cytochrome c did not change during its release from

the mitochondria.23

DNA damage in the cytotoxic process induced by silver

nanoparticles was shown to be induced by reactive oxygen

species (ROS).196 A similar, ROS-mediated toxicological response

was found for SWCNTs.61,197

Fig. 9 Investigation of the intraendosomal pH value. (A) SERS spectra of para-mercaptobenzoic acid (pMBA) coated gold nanoparticles at different

pH values. (B, C) The pH nanosensor is used to monitor the pH value inside live cells. Mouse fibroblast cells were incubated with pMBA gold

nanoparticles (pMBA concentration 1 mM) for 5 hours. (B) Selected SERS spectra of the pH nanosensor in vitro. (C) Raman chemical maps of fibroblast

cells visualize the local distribution of the intracellular pH value (intensity ratio of the pMBA Raman bands at 1076 cm�1 to 1424 cm�1). An excitation

wavelength of 785 nm was applied in all experiments with a laser intensity on the sample of B8 � 104 W cm�2 and an acquisition time of 1 s.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5794 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

Multi-walled carbon nanotubes were found to exert moderate

toxicity on HepG2 (liver hepatocellular carcinoma) cells. Molecu-

lar information of cellular constituents was retrieved in the

presence of the strong Raman contributions of lipid-functionalized

multi-walled carbon nanotubes.173 By using the high-frequency

region of the Raman spectra of cells containing the nanotubes,

the nucleus, cytoplasm, and lipid bodies were identified by the

intensity ratio of bands assigned to C–H stretching vibrations

of CH2 and CH3 groups. The localization of the characteristic

nucleus spectrum did not match that of the oxidized carbon

nanotubes, indicating that the material was not located in

the nucleus.173

The degree of DNA condensation varies strongly over the

course of the cell cycle, affecting the Raman spectra of the

nucleus. Notingher et al. have reported cell cycle phase

dependent Raman spectra, with main spectral differences

between the different stages occurring at a few characteristic

frequencies (782, 788, and 1095 cm�1).198 Later, Raman

microscopic images of the distribution of condensed chromatin

in the nuclei, using intensities of DNA and protein signals, could

be used to map cells in different phases of mitosis.199 Raman

spectra of human osteosarcoma cells that were synchronized in

G0/G1, S, and G2/M phases of the cell cycle were analyzed

with multivariate statistics, and changes in the cell spectra

were analyzed as a function of cell cycle phase, with principal

component analysis suggesting a decrease in the relative lipid

contribution in the course from G0/G1 to G2/M.200 Also spectral

contributions of the RNA vary in different cell cycle phases.201

Epigenetic changes, displaying as chromatin re-organization,

were also observed recently in Raman tweezer experiments with

Jurkat cells.202 A closer observation of the cell cycle is very

helpful when delineating the toxicity of a material: in the case of

silver nanoparticles, a possible toxic action may involve the

disruption of the respiratory chain in the mitochondria. The

resulting production of ROS interrupts generation of adenosine

triphosphate, leading to DNA damage.152 In addition, direct

interaction of the silver nanoparticles with the DNA is thought to

arrest the cell in the G2/M phase.152

Several groups have studied the chemical changes that occur

in the nuclei upon cell death, which are far more drastic than

those found for different stages in the cell cycle. The Raman

spectra of dead A549 lung epithelial cells indicated the break-

down of both phosphodiester bonds and DNA bases. In

particular, the intensity of a characteristic P–O–P band was

found to be extremely reduced.198,203 The fate of the nucleus and

its constituents during the apoptotic process can be sketched

using Raman data. In the earlier stage, the nucleus shrinks due to

the stress, as was shown in Raman imaging experiments by

Krafft et al.204 Later, the nucleus becomes fragmented, nucleo-

tide condensation and protein distribution in nuclear fragments

were mapped in apoptotic HeLa cells.205 In a very late apoptotic

stage, after fragmentation of the nucleus, when membrane

blebbing occurs at the surface of the cells, the nucleic acids show

up in the blebs.204 The apoptotic process was also investigated using

optical tweezer Raman spectroscopy.206 More recent imaging

of the time course of apoptosis indicates a high accumulation

of membrane phospholipids and highly unsaturated non-

membrane lipids in apoptotic cells, in addition to the DNA

condensation.207 Raman imaging is particularly useful to study

cell death, since the structural information that is obtained about

the nuclear components can be combined with the morphological

information on cellular structure. One example is the apoptotic

bodies, which are characteristic of the apoptotic process, but

absent in the case of necrosis. In necrotic cells, Raman spectro-

scopy indicated also a decrease in the amount of RNA, similar to

the observation in apoptosis, but a decrease in lipids.208

Raman spectroscopy on single cells, acting as sensors, has

become a tool for the detection of drug-induced cellular

changes, specifically drug-induced apoptosis.209,210 Recently, a

strategy of using Raman spectroscopy of single, isolated nuclei

was proposed for rapid and sensitive detection of cellular

changes in response to chemotherapeutic agents.210 These

results illustrate that Raman spectroscopy may as well become

a sensitive routine tool to assess nanotoxicity.

4.5 Nanoparticles in the nucleus

The transport of molecules between the cytoplasm and the

nucleus is known to occur through the nuclear pore complex

(NPC) and is mediated by multiple families of soluble trans-

port factors.211 All the transport receptors share the ability to

translocate across the NPC, probably through specific inter-

actions with components of the nuclear pore. Molecules larger

than 40 kDa must be transported by nucleo-cytoplasmic

shuttling receptors, recognizing their cargo in the cytoplasm

and then uploading it through the NPC to target the nucleus,

by signal- and energy-dependent mechanisms.212 The best

characterized nuclear import pathway involves cargo proteins,

comprising a basic amino acid-rich nuclear localization signal

(NLS).213–215 In order to enter the nucleus, also most nano-

particles will have to pass the NPC.

Several studies have succeeded in targeting the cellular

nucleus with gold nanoparticles, using different strategies.216–218

As the pioneering example, Feldherr and co-workers had

proven the existence and size exclusion of the NPC using

functionalized gold nanoparticles with oocyte nuclei.219 Later,

they demonstrated that the upper limit for nuclear import in

3T3 fibroblast cells is approximately 25 nm.220 A peptide

derived from the HIV Tat transactivator protein has been found

to act as an efficient molecule to incorporate gold NPs into

human fibroblasts and to translocate them into the nucleus.218

Oyelere and co-workers reported nuclear targeting with gold

nanorods carrying thiolated NLS peptide based on light

scattering in dark field microscopy.106 They also conducted

Raman experiments and found SERS spectral contributions

from the functional molecules on the nanorods.106 A recent

study using so-called cell-penetrating peptides reported that

the ultimate intracellular destination determined by the display of

the cell-penetrating peptide and further by the diameter of the

nanoparticles.90 The smallest 2.4 nm nanoparticles were found to

localize in the nucleus, while intermediate 5.5 and 8.2 nm

particles were partially delivered into the cytoplasm, showing a

perinuclear fate.90 Also silver nanoparticles have been targeted to

the nucleus using NLS peptides, and a cytotoxic response has

been monitored.221

In contrast, nuclear localization only rarely occurs in the

absence of localization signals. Ultrastructural studies using

quantitative analysis of transmission electron micrographs

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5795

have addressed the cellular uptake and intracellular trafficking of

15 nm gold nanoparticles, either plain or coated with PEG, exposed

to A549 lung epithelial cells. The data revealed a significant,

nonrandom intracellular distribution for both nanoparticle

types. No particles are observed in the nucleus, mitochondria,

endoplasmic reticulum, or Golgi.89 A study on uptake of silver

nanoparticles in human mesenchymal stem cells222 with light

microscopy reports similar findings: typically, silver agglomerates

were located in the perinuclear region, as determined by light

microscopy. Specific staining of cellular structures using fluorescent

probes showed that the silver nanoparticles occurred mainly within

endo-lysosomal structures, not in the cell nucleus, endoplasmic

reticulum or Golgi complex. In inhibition experiments, selectively

blocking the process of clathrin-dependent endocytosis indicated

that the nanoparticles were taken up primarily by clathrin-

dependent endocytosis and macropinocytosis.222

A few studies have reported SERS experiments with gold and

silver nanoparticles carrying nuclear targeting sequences in cul-

tured cells.106,168,223 Silver nanoparticles co-functionalized with

the TAT peptide showed greatly enhanced cellular uptake,

however without clear evidence for being taken up into the

nucleus.168 Dark field surface plasmon resonance scattering

images were used to demonstrate that NLS functionalized gold

nanorods are located in both the cytoplasm and nucleus of two

cell lines, an epithelial cell line and an oral cancer cell line.106 The

group reported SERS spectral contributions from nucleic acid

and protein components, which they assigned to the molecular

constituents of the nucleus.106 So far, ultrastructural evidence of

the nucleus being the exclusive origin of the nucleic acid and

protein signals in the SERS data has been lacking, and providing

such evidence will remain a great challenge. The probe volume

in a typical light microscope (usually on the order of tens

of femtoliters) employed in a Raman study always includes

cytoplasmic, perinuclear regions, where endocytotic vesicles with

extranuclear nanoparticles will accumulate. Due to the high

density of endosomes in this region, a very small portion of

cytoplasmic volume will be sufficient to generate high SERS

signals from the perinuclear region. As a second challenge, many

signals in a SERS spectrum that have been attributed to nucleic

acids and nuclear proteins were also found in experiments

where nuclear localization can be excluded based on the absence

of nuclear localization signals and/or large sizes of nano-

particles,116,165 and confirmation of a lack of nuclear localization

by electron microscopy.24 Especially in the late stage endolyso-

somes, nucleotides, DNA bases, and protein components can be

found in the proximity of the gold or silver nanoparticles. As

another aspect, nuclear integrity should be monitored carefully,

especially in experimental settings that could provoke pre-

apoptotic scenarios, such as incubation with silver nanoparticles,

or heating effects due to high plasmon absorption.224 The high

density in perinuclear late endosomes also requires very sophis-

ticated light microscopic approaches, or ideally high-resolution

ultrastructural microscopy experiments to ensure discrimination

between intranuclear and extranuclear nanoparticles.

5 Conclusions

To conclude, we have reviewed here work that concerns Raman

microspectroscopic studies of biological cells and tissues that

might be relevant for further investigations on the interaction

of nanomaterials with biological systems. The work discussed

here illustrates that a major strength of Raman microscopy

lies in the ability to map cellular chemistry, including the

possibility to monitor changes in living systems over time. In

principle, both, the nanomaterial and the biological structure

can be characterized in such a Raman experiment.

The Raman information can be obtained at different scales,

on whole organisms and tissues at the macroscopic scale, at

the microscopic, cellular level, and at the level of the inter-

action of biomolecules and cellular architecture with the

nanomaterials. Regarding the influence of the nanomaterials,

these different scales result in different levels of understanding.

A major challenge in further research will be the connection

between the different scales in different Raman experiments.

To a certain extent, this can be envisioned by designing

experiments which enable the observation of processes at both

the micro- and the nanoscale, when the structure of certain

molecules can be probed selectively, e.g., in some resonant

Raman microscopic experiments, or in SERS probing of

molecules adsorbing selectively at the nanoparticle surface.

Similarly, improved imaging technologies such as SORS

and hybrid Raman/fluorescence microscopies will need to be

employed to understand interaction of individual cells at the

tissue level, enabling the connection of cell physiological with

systemic, that is, microscopic with macroscopic information.

For the majority of molecular parameters that have to be

elucidated though, matching the information from different

Raman experiments will be the crucial task. An important

point here is the combination of experiments using relevant

models, so that macro- with microscale, and microscale with

nanoscale information can be connected. Specifically, suitable

cellular in vitro models that could mimic the situation in a

specific tissue should be used for Raman microscopy studies

on cells. This approach is obvious and becoming common

practice in toxicology research as well. A more complex task is

the choice of model at the (nanoscale) level of structure

investigations that are relevant for the situation of the inter-

action in the cell or tissue, including the nanomaterial. To this

end, more input will be needed from other biophysics and

molecular biology research. In order to successfully connect

knowledge on molecular structure and dynamics of cellular

molecules with that on biochemical and physiological

responses of cells or even of tissues, results from Raman

spectroscopy have to be combined with those of other inves-

tigations. For example, ultrastructural microscopy will reveal

complementary information on the interaction of nano-

particles with the cellular architecture, and cytotoxicology

investigations may provide cell physiological parameters

that could be combined with Raman information on the

biochemical composition of cellular organelles.

Acknowledgements

We would like to thank Betty Jobs for help with sorting

the literature. This work was supported in part by ERC

Grant 259432 (MULTIBIOPHOT) and funding from BAM

Innovationsoffensive (NANOTOX 1).

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5796 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

Notes and references

1 P. Matousek, Chem. Soc. Rev., 2007, 36, 1292–1304.2 A. S. Haka, K. E. Shafer-Peltier, M. Fitzmaurice, J. Crowe,R. R. Dasari and M. S. Feld, Proc. Natl. Acad. Sci. U. S. A.,2005, 102, 12371–12376.

3 K. Maquelin, C. Kirschner, L. P. Choo-Smith, N. van den Braak,H. P. Endtz, D. Naumann and G. J. Puppels, J. Microbiol.Methods, 2002, 51, 255–271.

4 J. Kneipp, H. Kneipp and K. Kneipp, Proc. Natl. Acad. Sci.U. S. A., 2006, 103, 17149–17153.

5 D. Serban, J. Benevides and G. Thomas Jr., Biochemistry, 2002,41, 847–853.

6 R. D. JiJi, G. Balakrishnan, Y. Hu and T. G. Spiro, Biochemistry,2006, 45, 34–41.

7 E. M. Jones, G. Balakrishnan and T. G. Spiro, J. Am. Chem. Soc.,2012, 134, 3461–3471.

8 S. A. Oladepo, K. Xiong, Z. Hong and S. A. Asher, J. Phys.Chem. Lett., 2011, 2, 334–344.

9 D. A. Heller, S. Baik, T. E. Eurell and M. S. Strano, Adv. Mater.,2005, 17, 2793–2799.

10 H. Hong, T. Gao and W. B. Cai, Nano Today, 2009, 4, 252–261.11 X. M. Qian, X. H. Peng, D. O. Ansari, Q. Yin-Goen, G. Z. Chen,

D. M. Shin, L. Yang, A. N. Young, M. D. Wang and S. M. Nie,Nat. Biotechnol., 2008, 26, 83–90.

12 S. Keren, C. Zavaleta, Z. Cheng, A. de la Zerda, O. Gheysens andS. S. Gambhir, Proc. Natl. Acad. Sci. U. S. A., 2008, 105,5844–5849.

13 T. Chernenko, C. Matthaus, L. Milane, L. Quintero, M. Amijiand M. Diem, ACS Nano, 2009, 3, 3552–3559.

14 Z. Liu, W. B. Cai, L. N. He, N. Nakayama, K. Chen, X. M. Sun,X. Y. Chen and H. J. Dai, Nat. Nanotechnol., 2007, 2, 47–52.

15 J. Qian, L. Jiang, F. H. Cai, D. Wang and S. L. He, Biomaterials,2011, 32, 1601–1610.

16 J. Kneipp, H. Kneipp and K. Kneipp, Chem. Soc. Rev., 2008, 37,1052–1060.

17 I. Chourpa, F. H. Lei, P. Dubois, M.Manfait andG. D. Sockalingum,Chem. Soc. Rev., 2008, 37, 993–1000.

18 H. J. Van Manen, N. Uzunbajakava, R. Van Bruggen, D. Roosand C. Otto, J. Am. Chem. Soc., 2003, 125, 12112–12113.

19 P. Candeloro, L. Tirinato, N. Malara, A. Fregola, E. Casals,V. Puntes, G. Perozziello, F. Gentile, M. L. Coluccio, G. Das,C. Liberale, F. De Angelis and E. Di Fabrizio, Analyst (Cambridge,U. K.), 2011, 136, 4402–4408.

20 P. Knief, C. Clarke, E. Herzog, M. Davoren, F. M. Lyng,A. D. Meade and H. J. Byrne, Analyst (Cambridge, U. K.), 2009,134, 1182–1191.

21 W. L. Peticolas, T. W. Patapoff, G. A. Thomas, J. Postlewait andJ. W. Powell, J. Raman Spectrosc., 1996, 27, 571–578.

22 C. Matthaus, T. Chernenko, J. A. Newmark, C. M. Warner andM. Diem, Biophys. J., 2007, 93, 668–673.

23 M. Okada, N. I. Smith, A. F. Palonpon, H. Endo, S. Kawata,M. Sodeoka and K. Fujita, Proc. Natl. Acad. Sci. U. S. A., 2012,109, 28–32.

24 J. Kneipp, H. Kneipp, M. McLaughlin, D. Brown andK. Kneipp, Nano Lett., 2006, 6, 2225–2231.

25 E. C. Faria and P. Gardner, Methods Mol. Biol. (Clifton, N.J.),2012, 853, 151–167.

26 R. D. Snook, T. J. Harvey, E. C. Faria and P. Gardner, Integr.Biol., 2009, 1, 43–52.

27 P. R. T. Jess, V. Garces-Chavez, D. Smith, M. Mazilu,L. Paterson, A. Riches, C. S. Herrington, W. Sibbett andK. Dholakia, Opt. Express, 2006, 14, 5779–5791.

28 K. Ramser, K. Logg, M. Goksor, J. Enger, M. Kall andD. Hanstorp, J. Biomed. Opt., 2004, 9, 593–600.

29 C. G. Xie, M. A. Dinno and Y. Q. Li, Opt. Lett., 2002, 27,249–251.

30 H. Tang, H. Yao, G. Wang, Y. Wang, Y.-Q. Li and M. Feng,Opt. Express, 2007, 15, 12708–12716.

31 U. P. Agarwal and R. H. Atalla, Planta, 1986, 169, 325–332.32 R. H. Atalla and U. P. Agarwal, Science, 1985, 227, 636–638.33 L. Q. Chu, R. Masyuko, J. V. Sweedler and P.W. Bohn, Bioresour.

Technol., 2010, 101, 4919–4925.34 S. Richter, J. Mussig and N. Gierlinger, Planta, 2011, 233,

763–772.

35 M. Schmidt, A. M. Schwartzberg, A. Carroll, A. Chaibang,P. D. Adams and P. J. Schuck, Biochem. Biophys. Res. Commun.,2010, 395, 521–523.

36 Y. Morikawa, A. Yoshinaga, H. Kamitakahara, M. Wada andK. Takabe, Holzforschung, 2010, 64, 61–67.

37 P. Robert, F. Jamme, C. Barron, B. Bouchet, L. Saulnier,P. Dumas and F. Guillon, Planta, 2011, 233, 393–406.

38 H. Wu, J. V. Volponi, A. E. Oliver, A. N. Parikh, B. A. Simmonsand S. Singh, Proc. Natl. Acad. Sci. U. S. A., 2011, 108,3809–3814.

39 P. Heraud, J. Beardall, D. McNaughton and B. R. Wood, FEMSMicrobiol. Lett., 2007, 275, 24–30.

40 B. R. Wood, P. Heraud, S. Stojkovic, D. Morrison, J. Beardalland D. McNaughton, Anal. Chem. (Washington, DC, U. S.),2005, 77, 4955–4961.

41 O. Samek, A. Jonas, Z. Pilat, P. Zemanek, L. Nedbal, J. Triska,P. Kotas and M. Trtilek, Sensors, 2010, 10, 8635–8651.

42 F. Schulte, U. Panne and J. Kneipp, J. Biophotonics, 2010, 3,542–547.

43 S. Busch, K. Schmitt, C. Erhardt and T. Speck, J. RamanSpectrosc., 2010, 41, 490–497.

44 C. M. Muntean, N. Leopold, A. Halmagyi and S. Valimareanu,J. Raman Spectrosc., 2010, 42, 844–850.

45 A. Shen, J. Guo, W. Xie, M. Sun, R. Richards and J. Hu,J. Raman Spectrosc., 2011, 42, 879–884.

46 R. A. Alvarez-Puebla and L. M. Liz-Marzan, Energy Environ.Sci., 2010, 3, 1011–1017.

47 L. Zeiri, J. Raman Spectrosc., 2007, 38, 950–955.48 M. Alimohammadi, Y. Xu, D. Y. Wang, A. S. Biris and

M. V. Khodakovskaya, Nanotechnology, 2011, 22.49 E. M. Jorgensen and S. E. Mango, Nat. Rev. Genet., 2002, 3,

356–369.50 S. Charan, F.-C. Chien, N. Singh, C.-W. Kuo and P. Chen,

Chem.–Eur. J., 2011, 17, 5165–5170.51 C. Onogi and H.-O. Hamaguchi, J. Phys. Chem. B, 2009, 113,

10942–10945.52 C. Onogi, H. Torii and H.-o. Hamaguchi, Chem. Lett., 2009, 38,

898–899.53 A. Sujith, T. Itoh, H. Abe, A. A. Anas, K. Yoshida, V. Biju and

M. Ishikawa, Appl. Phys. Lett., 2008, 92.54 A. Sujith, T. Itoh, H. Abe, K.-i. Yoshida, M. S. Kiran, V. Biju

and M. Ishikawa, Anal. Bioanal. Chem., 2009, 394, 1803–1809.55 R. F. Fakhrullin, A. I. Zamaleeva, M. V.Morozov, D. I. Tazetdinova,

F. K. Alimova, A. K. Hilmutdinov, R. I. Zhdanov,M.Kahraman andM. Culha, Langmuir, 2009, 25, 4628–4634.

56 D. Pantarotto, R. Singh, D. McCarthy, M. Erhardt, J. P. Briand,M. Prato, K. Kostarelos and A. Bianco, Angew. Chem., Int. Ed.,2004, 43, 5242–5246.

57 D. Pantarotto, J. P. Briand, M. Prato and A. Bianco, Chem.Commun., 2004, 16–17.

58 A. M. Rao, E. Richter, S. Bandow, B. Chase, P. C. Eklund,K. A. Williams, S. Fang, K. R. Subbaswamy, M. Menon,A. Thess, R. E. Smalley, G. Dresselhaus and M. S. Dresselhaus,Science, 1997, 275, 187–191.

59 S. T. Yang, X. Wang, G. Jia, Y. Q. Gu, T. C. Wang, H. Y. Nie,C. C. Ge, H. F. Wang and Y. F. Liu, Toxicol. Lett., 2008, 181,182–189.

60 M. L. Schipper, N. Nakayama-Ratchford, C. R. Davis, N. W.S. Kam, P. Chu, Z. Liu, X. M. Sun, H. J. Dai and S. S. Gambhir,Nat. Nanotechnol., 2008, 3, 216–221.

61 Y. B. Zhang, Y. Xu, Z. G. Li, T. Chen, S. M. Lantz,P. C. Howard, M. G. Paule, W. Slikker, F. Watanabe,T. Mustafa, A. S. Biris and S. F. Ali, ACS Nano, 2011, 5,7020–7033.

62 C. Zavaleta, A. de la Zerda, Z. Liu, S. Keren, Z. Cheng,M. Schipper, X. Chen, H. Dai and S. S. Gambhir, Nano Lett.,2008, 8, 2800–2805.

63 A. S. Biris, E. I. Galanzha, Z. Li, M. Mahmood, Y. Xu andV. P. Zharov, J. Biomed. Opt., 2009, 14.

64 Z. Liu, C. Davis, W. B. Cai, L. He, X. Y. Chen and H. J. Dai,Proc. Natl. Acad. Sci. U. S. A, 2008, 105, 1410–1415.

65 A. S. Thakor, R. Luong, R. Paulmurugan, F. I. Lin, P. Kempen,C. Zavaleta, P. Chu, T. F. Massoud, R. Sinclair andS. S. Gambhir, Sci. Transl. Med., 2011, 3.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5797

66 C. L. Zavaleta, B. R. Smith, I. Walton, W. Doering, G. Davis,B. Shojaei, M. J. Natan and S. S. Gambhir, Proc. Natl. Acad. Sci.U. S. A., 2009, 106, 13511–13516.

67 G. von Maltzahn, A. Centrone, J.-H. Park, R. Ramanathan,M. J. Sailor, T. A. Hatton and S. N. Bhatia, Adv. Mater., 2009,21, 3175–3180.

68 J.-H. Park, G. von Maltzahn, L. L. Ong, A. Centrone,T. A. Hatton, E. Ruoslahti, S. N. Bhatia and M. J. Sailor, Adv.Mater., 2010, 22, 880–885.

69 A. J. Hobro and B. Lendl, Vib. Spectrosc., 2011, 57, 213–219.70 O. Burkacky, A. Zumbusch, C. Brackmann and A. Enejder, Opt.

Lett., 2006, 31, 3656–3658.71 C. W. Freudiger, W. Min, G. R. Holtom, B. Xu, M. Dantus and

X. S. Xie, Nat. Photonics, 2011, 5, 103–109.72 J. P. R. Day, G. Rago, K. F. Domke, K. P. Velikov andM. Bonn,

J. Am. Chem. Soc., 2010, 132, 8433–8439.73 Y. L. Wang, J. L. Seebald, D. P. Szeto and J. Irudayaraj, ACS

Nano, 2010, 4, 4039–4053.74 P. J. Caspers, G. W. Lucassen, E. A. Carter, H. A. Bruining and

G. J. Puppels, J. Invest. Dermatol., 2001, 116, 434–442.75 P. Matousek, M. D. Morris, N. Everall, I. P. Clark, M. Towrie,

E. Draper, A. Goodship and A. W. Parker, Appl. Spectrosc.,2005, 59, 1485–1492.

76 A. Mahadevan-Jansen, M. F. Mitchell, N. Ramanujam,A. Malpica, S. Thomsen, U. Utzinger and R. Richards-Kortum,Photochem. Photobiol., 1998, 68, 123–132.

77 P. Matousek, E. R. C. Draper, A. E. Goodship, I. P. Clark,K. L. Ronayne and A. W. Parker, Appl. Spectrosc., 2006, 60,758–763.

78 M. V. Schulmerich, J. H. Cole, J. M. Kreider, F. Esmonde-White,K. A. Dooley, S. A. Goldstein and M. D. Morris, Appl. Spectrosc.,2009, 63, 286–295.

79 M. V. Schulmerich, J. H. Cole, K. A. Dooley, M. D. Morris,J. M. Kreider, S. A. Goldstein, S. Srinivasan and B. W. Pogue,J. Biomed. Opt., 2008, 13.

80 N. Stone, K. Faulds, D. Graham and P. Matousek, Anal. Chem.,2010, 82, 3969–3973.

81 N. Stone, M. Kerssens, G. R. Lloyd, K. Faulds, D. Graham andP. Matousek, Chem. Sci., 2011, 2, 776–780.

82 J. M. Yuen, N. C. Shah, J. T. Walsh Jr, M. R. Glucksberg andR. P. Van Duyne, Anal. Chem., 2010, 82, 8382–8385.

83 D. Drescher, G. Orts-Gil, G. Laube, K. Natte, R. W. Veh,W. Osterle and J. Kneipp, Anal. Bioanal. Chem., 2011, 400,1367–1373.

84 J. Lin, R. Chen, S. Feng, Y. Li, Z. Huang, S. Xie, Y. Yu,M. Cheng and H. Zeng, Biosens. Bioelectron., 2009, 25, 388–394.

85 Y. Yu, J.-Q. Lin, H. Huang, Y.-P. Chen, S.-Y. Feng, Y. Su,J.-S. Chen and R. Chen, Prog. Biochem. Biophys., 2011, 38,961–966.

86 Y. Yu, J. Lin, Y. Wu, S. Feng, Y. Li, Z. Huang, R. Chen andH. Zeng, Spectrosc. Int. J., 2011, 25, 13–21.

87 P. Nativo, I. A. Prior and M. Brust, ACS Nano, 2008, 2,1639–1644.

88 D. B. Chithrani, Mol. Membr. Biol., 2010, 27, 299–311.89 C. Brandenberger, C. Muhlfeld, Z. Ali, A. G. Lenz, O. Schmid,

W. J. Parak, P. Gehr and B. Rothen-Rutishauser, Small, 2010, 6,1669–1678.

90 E. Oh, J. B. Delehanty, K. E. Sapsford, K. Susumu, R. Goswami,J. B. Blanco-Canosa, P. E. Dawson, J. Granek, M. Shoff,Q. Zhang, P. L. Goering, A. Huston and I. L. Medintz, ACSNano, 2011, 5, 6434–6448.

91 B. D. Chithrani, A. A. Ghazani and W. C. W. Chan, Nano Lett.,2006, 6, 662–668.

92 H. Jin, D. A. Heller, R. Sharma and M. S. Strano, ACS Nano,2009, 3, 149–158.

93 A. E. Nel, L. Madler, D. Velegol, T. Xia, E. M. V. Hoek,P. Somasundaran, F. Klaessig, V. Castranova andM. Thompson, Nat. Mater., 2009, 8, 543–557.

94 O. Lunov, T. Syrovets, C. Loos, J. Beil, M. Delecher, K. Tron,G. U. Nienhaus, A. Musyanovych, V. Mailander, K. Landfesterand T. Simmet, ACS Nano, 2011, 5, 1657–1669.

95 K. Kostarelos, L. Lacerda, G. Pastorin, W. Wu, S. Wieckowski,J. Luangsivilay, S. Godefroy, D. Pantarotto, J.-P. Briand,S. Muller, M. Prato and A. Bianco, Nat. Nanotechnol., 2007, 2,108–113.

96 I. Canton and G. Battaglia, Chem. Soc. Rev., 2012, 41,2718–2739.

97 S. D. Conner and S. L. Schmid, Nature, 2003, 422, 37–44.98 W. Li, C. Y. Chen, C. Ye, T. T. Wei, Y. L. Zhao, F. Lao, Z. Chen,

H. Meng, Y. X. Gao, H. Yuan, G. M. Xing, F. Zhao, Z. F. Chai,X. J. Zhang, F. Y. Yang, D. Han, X. H. Tang and Y. G. Zhang,Nanotechnology, 2008, 19.

99 J. Rejman, V. Oberle, I. S. Zuhorn and D. Hoekstra, Biochem. J,2004, 377, 159–169.

100 A. A. van Apeldoorn, H. J. van Manen, J. M. Bezemer, J. D. deBruijn, C. A. van Blitterswijk and C. Otto, J. Am. Chem. Soc.,2004, 126, 13226–13227.

101 L. Tetard, A. Passian, K. T. Venmar, R. M. Lynch, B. H. Voy,G. Shekhawat, V. P. Dravid and T. Thundat, Nat. Nanotechnol.,2008, 3, 501–505.

102 J. Ando, K. Fujita, N. I. Smith and S. Kawata, Nano Lett., 2011,11, 5344–5348.

103 K. Fujita, S. Ishitobi, K. Hamada, N. I. Smith, A. Taguchi,Y. Inouye and S. Kawata, J. Biomed. Opt., 2009, 14.

104 J. Kneipp, H. Kneipp, B. Wittig and K. Kneipp,Nano Lett., 2007,7, 2819–2823.

105 J. Kneipp, H. Kneipp, B. Wittig and K. Kneipp, J. Phys. Chem.C, 2010, 7421–7426.

106 A. K. Oyelere, P. C. Chen, X. Huang, I. H. El-Sayed andM. A. El-Sayed, Bioconjugate Chem., 2007, 18, 1490–1497.

107 M. S. Dresselhaus, G. Dresselhaus, A. Jorio, A. G. Souza,M. A. Pimenta and R. Saito, Acc. Chem. Res., 2002, 35,1070–1078.

108 H.-J. van Manen, Y. M. Kraan, D. Roos and C. Otto, Proc. Natl.Acad. Sci. U. S. A., 2005, 102, 10159–10164.

109 R. J. Stokes, F. McKenzie, E. McFarlane, A. Ricketts, L. Tetley,K. Faulds, J. Alexander and D. Graham, Analyst, 2009, 134,170–175.

110 P. Chen, Y. Chen, L. Su, A. Shen, J. Hu, X. Wang and J. Hu,J. Mater. Chem., 2012, 22, 631–635.

111 C. L. Haynes, A. D. McFarland and R. P. Van Duyne, Anal.Chem., 2005, 77, 338A–346A.

112 Z. Liu, M. Winters, M. Holodniy and H. Dai, Angew. Chem., Int.Ed., 2007, 46, 2023–2027.

113 J. Kneipp, H. Kneipp, W. L. Rice and K. Kneipp, Anal. Chem.,2005, 77, 2381–2385.

114 J. Kneipp, H. Kneipp, A. Rajaduraj, R. W. Redmond andK. Kneipp, J. Raman Spectrosc., 2009, 40, 1–5.

115 X. Tan, Z. Wang, J. Yang, C. Song, R. Zhang and Y. Cui,Nanotechnology, 2009, 20.

116 H.-W. Tang, X. B. Yang, J. Kirkham and D. A. Smith, Anal.Chem., 2007, 79, 3646–3653.

117 C. D. Syme, N. M. S. Sirimuthu, S. L. Faley and J. M. Cooper,Chem. Commun., 2010, 7921–7923.

118 J. Xie, Q. Zhang, J. Y. Lee and D. I. C. Wang, ACS Nano, 2008,2, 2473–2480.

119 M. K. Gregas, F. Yan, J. Scaffidi, H. N. Wang and T. Vo-Dinh,Nanomedicine (Philadelphia, PA, U. S.), 2011, 7, 115–122.

120 A. Matschulat, D. Drescher and J. Kneipp, ACS Nano, 2010, 4,3259–3269.

121 N. M. S. Sirimuthu, C. D. Syme and J. M. Cooper, Chem.Commun., 2011, 4099–4101.

122 C. E. Talley, L. Jusinski, C. W. Hollars, S. M. Lane and T. Huser,Anal. Chem., 2004, 76, 7064–7068.

123 J. H. Park, J. Park, U. Dembereldorj, K. Cho, K. Lee, S. I. Yang,S. Y. Lee and S. W. Joo, Anal. Bioanal. Chem., 2011, 401,1631–1639.

124 Z. Y. Wang, H. Wu, C. L. Wang, S. H. Xu and Y. P. Cui,J. Mater. Chem., 2011, 21, 4307–4313.

125 A. Shamsaie, J. Heim, A. A. Yanik and J. Irudayaraj, Chem.Phys. Lett., 2008, 461, 131–135.

126 N. M. S. Sirimuthu, C. D. Syme and J. M. Cooper, Anal. Chem.,2010, 82, 7369–7373.

127 S. Schlucker, B. Kustner, A. Punge, R. Bonfig, A. Marx andP. Strobel, J. Raman Spectrosc., 2006, 37, 719–721.

128 Q. Hu, L.-L. Tay, M. Noestheden and J. P. Pezacki, J. Am. Chem.Soc., 2007, 129, 14–15.

129 A. L. Koh, C. M. Shachaf, S. Elchuri, G. P. Nolan andR. Sinclair, Ultramicroscopy, 2008, 109, 111–121.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

5798 Chem. Soc. Rev., 2012, 41, 5780–5799 This journal is c The Royal Society of Chemistry 2012

130 B. R. Lutz, C. E. Dentinger, L. N. Nguyen, L. Sun, J. Zhang,A. N. Allen, S. Chan and B. S. Knudsen, ACS Nano, 2008, 2,2306–2314.

131 K. Hering, D. Cialla, K. Ackermann, T. Doerfer, R. Moeller,H. Schneidewind, R. Mattheis, W. Fritzsche, P. Roesch andJ. Popp, Anal. Bioanal. Chem., 2008, 390, 113–124.

132 L. Sun, K.-B. Sung, C. Dentinger, B. Lutz, L. Nguyen, J. Zhang,H. Qin, M. Yamakawa, M. Cao, Y. Lu, A. J. Chmura, J. Zhu,X. Su, A. A. Berlin, S. Chan and B. Knudsen, Nano Lett., 2007, 7,351–356.

133 D. C. Kennedy, C. S. McKay, L.-l. Tay, Y. Rouleau andJ. P. Pezacki, Chem. Commun., 2011, 3156–3158.

134 D. C. Kennedy, K. A. Hoop, L.-L. Tay and J. P. Pezacki,Nanoscale, 2010, 2, 1413–1416.

135 Z. Liu, S. Tabakman, S. Sherlock, X. L. Li, Z. Chen, K. L. Jiang,S. S. Fan and H. J. Dai, Nano Res., 2010, 3, 222–233.

136 H. J. van Manen, A. A. van Apeldoorn, R. Verrijk, C. A. vanBlitterswijk and C. Otto, Int. J. Nanomed., 2007, 2, 241–252.

137 S. Reich, C. Thomsen and P. Ordejon, Phys. Rev. B: Condens.Matter Mater Phys., 2002, 65.

138 D. A. Heller, P. W. Barone, J. P. Swanson, R. M. Mayrhofer andM. S. Strano, J. Phys. Chem. B, 2004, 108, 6905–6909.

139 M. S. Strano, M. Zheng, A. Jagota, G. B. Onoa, D. A. Heller,P. W. Barone and M. L. Usrey, Nano Lett., 2004, 4, 543–550.

140 C. G. Blatchford, J. R. Campbell and J. A. Creighton, Surf. Sci.,1982, 120, 435–455.

141 M. Kerker, O. Siiman and D. S. Wang, J. Phys. Chem., 1984, 88,3168–3170.

142 P. Hildebrandt, S. Keller, A. Hoffmann, F. Vanhecke andB. Schrader, J. Raman Spectrosc., 1993, 24, 791–796.

143 E. Hao and G. C. Schatz, J. Chem. Phys., 2004, 120, 357–366.144 M. Moskovits, J. Raman Spectrosc., 2005, 36, 485–496.145 K. Kneipp, H. Kneipp and J. Kneipp, Acc. Chem. Res., 2006, 39,

443–450.146 A. Otto, I. Mrozek, H. Grabhorn and W. Akemann, J. Phys.:

Condens. Matter, 1992, 4, 1143–1212.147 V. Joseph, A. Matschulat, J. Polte, S. Rolf, F. Emmerling and

J. Kneipp, J. Raman Spectrosc., 2011, 42, 1736–1742.148 M. I. Stockman, V. M. Shalaev, M. Moskovits, R. Botet and

T. F. George, Phys. Rev. B: Condens. Matter Mater. Phys., 1992,46, 2821.

149 H. X. Xu, J. Aizpurua, M. Kall and P. Apell, Phys. Rev. E:Stat. Phys., Plasmas, Fluids, Relat. Interdiscip. Top., 2000, 62,4318–4324.

150 K. R. Li, M. I. Stockman and D. J. Bergman, Phys. Rev. Lett.,2003, 91, 227402.

151 P. J. G. Goulet, D. S. dos Santos, R. A. Alvarez-Puebla,O. N. Oliveira and R. F. Aroca, Langmuir, 2005, 21, 5576–5581.

152 P. V. Asharani, G. L. K. Mun, M. P. Hande and S. Valiyaveettil,ACS Nano, 2009, 3, 279–290.

153 S. W. P. Wijnhoven, W. J. G. M. Peijnenburg, C. A. Herberts,W. I. Hagens, A. G. Oomen, E. H. W. Heugens, B. Roszek,J. Bisschops, I. Gosens, D. Van de Meent, S. Dekkers, W. H. DeJong, M. Van Zijverden, A. J. A. M. Sips and R. E. Geertsma,Nanotoxicology, 2009, 3, 109–U178.

154 M. C. Stensberg, Q. S. Wei, E. S. McLamore, D. M. Porterfield,A. Wei and M. S. Sepulveda, Nanomedicine (Philadelphia, PA,U. S.), 2011, 6, 879–898.

155 H. J. Johnston, G. Hutchison, F. M. Christensen, S. Peters,S. Hankin and V. Stone, Crit. Rev. Toxicol., 2010, 40, 328–346.

156 G. Buscarino, V. Ardizzone, G. Vaccaro, S. Agnello andF. M. Gelardi, J. Appl. Phys., 2010, 108.

157 G. J. Puppels, H. S. P. Garritsen, G. M. J. Segersnolten, F. F. M.Demul and J. Greve, Biophys. J., 1991, 60, 1046–1056.

158 N. M. Sijtsema, C. Otto, G. M. J. Segers-Nolten, A. J. Verhoevenand J. Greve, Biophys. J., 1998, 74, 3250–3255.

159 H. J. Van Manen and C. Otto, Nano Lett., 2007, 7, 1631–1636.160 W.-T. Chang, Y.-C. Yang, H.-H. Lu, I. L. Li and I. Liau, J. Am.

Chem. Soc., 2010, 132, 1744–1745.161 S. Sanchezcortes and J. V. Garciaramos, J. Mol. Struct., 1992,

274, 33–45.162 R. W. VanDyke, Am. J. Physiol., 1993, 265, C901–C917.163 M. Grabe and G. Oster, J. Gen. Physiol., 2001, 117, 329–344.164 R. W. Van Dyke, Hepatology, 2000, 32, 1357–1369.

165 K. Kneipp, A. S. Haka, H. Kneipp, K. Badizadegan, N. Yoshizawa,C. Boone, K. E. Shafer-Peltier, J. T. Motz, R. R. Dasari andM. S. Feld, Appl. Spectrosc., 2002, 56, 150–154.

166 J. Kneipp, H. Kneipp, B. Wittig and K. Kneipp, Nanomedicine(Philadelphia, PA, U. S.), 2010, 6, 214–226.

167 R. R. Sathuluri, H. Yoshikawa, E. Shimizu, M. Saito andE. Tamiya, PLoS One, 2011, 6.

168 M. K. Gregas, J. P. Scaffidi, B. Lauly and T. Vo-Dinh, Appl.Spectrosc., 2010, 64, 858–866.

169 B. Moody, C. M. Haslauer, E. Kirk, A. Kannan, E. G. Loboaand G. S. McCarty, Appl. Spectrosc., 2010, 64, 1227–1233.

170 W. Xie, C. Herrmann, K. Koempe, M. Haase and S. Schluecker,J. Am. Chem. Soc., 2011, 133, 19302–19305.

171 V. Joseph, M. Gensler, S. Seifert, U. Gernert, J. P. Rabe andJ. Kneipp, J. Phys. Chem. C, 2012, 116, 6859–6865.

172 J. Dorney, F. Bonnier, A. Garcia, A. Casey, G. Chambers andH. J. Byrne, Analyst, 2011, 137, 1111–1119.

173 G. Romero, I. Estrela-Lopis, P. Castro-Hartmann, E. Rojas,I. Llarena, D. Sanz, E. Donath and S. E. Moya, Soft Matter,2011, 7, 6883–6890.

174 C. Onogi, M.Motoyama andH.-o. Hamaguchi, J. Raman Spectrosc.,2008, 39, 555–556.

175 R. Boehme, D. Cialla, M. Richter, P. Roesch, J. Popp andV. Deckert, J. Biophotonics, 2010, 3, 455–461.

176 R. Boehme, M. Richter, D. Cialla, P. Roesch, V. Deckert andJ. Popp, J. Raman Spectrosc., 2009, 40, 1452–1457.

177 T. Schmid, B.-S. Yeo, G. Leong, J. Stadler and R. Zenobi,J. Raman Spectrosc., 2009, 40, 1392–1399.

178 R. F. Murphy, S. Powers and C. R. Cantor, J. Cell Biol., 1984, 98,1757–1762.

179 P. Montcourrier, P. H. Mangeat, C. Valembois, G. Salazar,A. Sahuquet, C. Duperray and H. Rochefort, J. Cell Sci., 1994,107, 2381–2391.

180 A. Michota and J. Bukowska, J. Raman Spectrosc., 2003, 34,21–25.

181 S. W. Bishnoi, C. J. Rozell, C. S. Levin, M. K. Gheith,B. R. Johnson, D. H. Johnson and N. J. Halas, Nano Lett.,2006, 6, 1687–1692.

182 R. A. Jensen, J. Sherin and S. R. Emory, Appl. Spectrosc., 2007,61, 832–838.

183 A. M. Schwartzberg, T. Y. Oshiro, J. Z. Zhang, T. Huser andC. E. Talley, Anal. Chem., 2006, 78, 4732–4736.

184 M. A. Ochsenkuhn, P. R. T. Jess, H. Stoquert, K. Dholakia andC. J. Campbell, ACS Nano, 2009, 3, 3613–3621.

185 J. P. Scaffidi, M. K. Gregas, V. Seewaldt and T. Vo-Dinh, Anal.Bioanal. Chem., 2009, 393, 1135–1141.

186 Z. Wang, A. Bonoiu, M. Samoc, Y. Cui and P. N. Prasad,Biosens. Bioelectron., 2008, 23, 886–891.

187 S. Zong, Z. Wang, J. Yang and Y. Cui, Anal. Chem., 2011, 83,4178–4183.

188 J. F. Poschet, J. A. Fazio, G. S. Timmins, W. Ornatowski,E. Perkett, M. Delgado and V. Deretic, EMBO Rep., 2006, 7,553–559.

189 M. Hara-Chikuma, Y. H. Wang, S. E. Guggino, W. B. Gugginoand A. S. Verkman, Biochem. Biophys. Res. Commun., 2005, 329,941–946.

190 N. Kokkonen, A. Rivinoja, A. Kauppila, M. Suokas,I. Kellokumpu and S. Kellokumpu, J. Biol. Chem., 2004, 279,39982–39988.

191 J. W. Chan, D. S. Taylor, T. Zwerdling, S. M. Lane, K. Ihara andT. Huser, Biophys. J., 2006, 90, 648–656.

192 F. Draux, P. Jeannesson, A. Beljebbar, A. Tfayli, N. Fourre,M. Manfait, J. Sule-Suso and G. D. Sockalingum, Analyst, 2009,134, 542–548.

193 K. Klein, A. M. Gigler, T. Aschenbrenne, R. Monetti, W. Bunk,F. Jamitzky, G. Morfill, R. W. Stark and J. Schlegel, Biophys. J.,2012, 102, 360–368.

194 A. V. Feofanov, A. I. Grichine, L. A. Shitova, T. A. Karmakova,R. I. Yakubovskaya, M. Egret-Charlier and P. Vigny, Biophys. J.,2000, 78, 499–512.

195 K. Meister, J. Niesel, U. Schatzschneider, N. Metzler-Nolte,D. A. Schmidt and M. Havenith, Angew. Chem., Int. Ed., 2010,49, 3310–3312.

196 R. Foldbjerg, D. A. Dang and H. Autrup, Arch. Toxicol., 2011,85, 743–750.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online

This journal is c The Royal Society of Chemistry 2012 Chem. Soc. Rev., 2012, 41, 5780–5799 5799

197 H. F. Cui, S. K. Vashist, K. Al-Rubeaan, J. H. T. Luong andF. S. Sheu, Chem. Res. Toxicol., 2010, 23, 1131–1147.

198 I. Notingher, S. Verrier, S. Haque, J. M. Polak and L. L. Hench,Biopolymers, 2003, 72, 230–240.

199 C. Matthaus, S. Boydston-White, M. Miljkovic, M. Romeo andM. Diem, Appl. Spectrosc., 2006, 60, 1–8.

200 R. J. Swain, G. Jell and M. A. Stevens, J. Cell. Biochem., 2008,104, 1427–1438.

201 V. V. Pully, A. Lenferink and C. Otto, Vib. Spectrosc., 2010, 53,12–18.

202 M. Poplineau, A. Trussardi-Regnier, T. Happillon, J. Dufer,M. Manfait, P. Bernard, O. Piot and F. Antonicelli, Epigenomics,2011, 3, 785–794.

203 S. Verrier, I. Notingher, J. M. Polak and L. L. Hench, Biopolymers,2004, 74, 157–162.

204 C. Krafft, T. Knetschke, R. H. W. Funk and R. Salzer, Anal.Chem., 2006, 78, 4424–4429.

205 N. Uzunbajakava, A. Lenferink, Y. Kraan, E. Volokhina, G. Vrensen,J. Greve and C. Otto, Biophys. J., 2003, 84, 3968–3981.

206 T. J. Moritz, D. S. Taylor, D. M. Krol, J. Fritch and J. W. Chan,Biomed. Opt. Express, 2010, 1, 1138–1147.

207 A. Zoladek, F. C. Pascut, P. Patel and I. Notingher, J. RamanSpectrosc., 2011, 42, 251–258.

208 N. Kunapareddy, J. P. Freyer and J. R. Mourant, J. Biomed.Opt., 2008, 13.

209 R. Buckmaster, F. Asphahani, M. Thein, J. Xu and M. Zhang,Analyst, 2009, 134, 1440–1446.

210 H.-H. Lin, Y.-C. Li, C.-H. Chang, C. Liu, A. L. Yu andC.-H. Chen, Anal. Chem., 2012, 84, 113–120.

211 D. Stoffler, K. N. Goldie, B. Feja and U. Aebi, J. Mol. Biol., 1999,287, 741–752.

212 D. Gorlich and U. Kutay, Annu. Rev. Cell Dev. Biol., 1999, 15,607–660.

213 W. Ritter, C. Plank, J. Lausier, C. Rudolph, D. Zink,D. Reinhardt and J. Rosenecker, J. Mol. Med., 2003, 81, 708–717.

214 I. Schmitt and L. Gerace, J. Biol. Chem., 2001, 276, 42355–42363.215 A. Lange, R. E. Mills, C. J. Lange, M. Stewart, S. E. Devine and

A. H. Corbett, J. Biol. Chem., 2007, 282, 5101–5105.216 A. G. Tkachenko, H. Xie, Y. L. Liu, D. Coleman, J. Ryan,

W. R. Glomm, M. K. Shipton, S. Franzen and D. L. Feldheim,Bioconjugate Chem., 2004, 15, 482–490.

217 A. G. Tkachenko, H. Xie, D. Coleman, W. Glomm, J. Ryan,M. F. Anderson, S. Franzen and D. L. Feldheim, J. Am. Chem.Soc., 2003, 125, 4700–4701.

218 J. M. de la Fuente and C. C. Berry, Bioconjugate Chem., 2005, 16,1176–1180.

219 C. Feldherr, E. Kallenbach and N. Schultz, J. Cell Biol., 1984, 99,2216–2222.

220 C. M. Feldherr, D. Akin and R. J. Cohen, J. Cell Sci., 2001, 114,4621–4627.

221 L. A. Austin, B. Kang, C.-W. Yen and M. A. El-Sayed, J. Am.Chem. Soc., 2011, 133, 17594–17597.

222 C. Greulich, J. Diendorf, T. Simon, G. Eggeler, M. Epple andM. Koller, Acta Biomater., 2011, 7, 347–354.

223 W. Xie, L. Wang, Y. Zhang, L. Su, A. Shen, J. Tan and J. Hu,Bioconjugate Chem., 2009, 20, 768–773.

224 R. A. Sperling, P. Rivera gil, F. Zhang, M. Zanella andW. J. Parak, Chem. Soc. Rev., 2008, 37, 1896–1908.

Dow

nloa

ded

by U

nive

rsity

of

Gla

sgow

Lib

rary

on

14 M

arch

201

3Pu

blis

hed

on 1

1 Ju

ly 2

012

on h

ttp://

pubs

.rsc

.org

| do

i:10.

1039

/C2C

S351

27G

View Article Online