Journal of Physics D: Applied Physics
J. Phys. D: Appl. Phys. 51 (2018) 443001 (42pp)

https://doi.org/10.1088/1361-6463/aad055

Topical Review

The 2018 correlative microscopy
techniques roadmap
Toshio Ando1 , Satya Prathyusha Bhamidimarri2, Niklas Brending3,
H Colin-York4 , Lucy Collinson5, Niels De Jonge6,7, P J de Pablo8,9 ,Elke Debroye10, Christian Eggeling4,11,12,33 , Christian Franck13,
Marco Fritzsche4,14, Hans Gerritsen15, Ben N G Giepmans16,
Kay Grunewald17,18,19, Johan Hofkens10, Jacob P Hoogenboom20,33 ,Kris P F Janssen10 , Rainer Kaufmann17,18,21, Judith Klumpermann22 ,Nyoman Kurniawan23, Jana Kusch24, Nalan Liv22 , Viha Parekh23,
Diana B Peckys25, Florian Rehfeldt26 , David C Reutens23,
Maarten B J Roeffaers27, Tim Salditt28, Iwan A T Schaap29,33 ,Ulrich S Schwarz30, Paul Verkade31, Michael W Vogel23, Richard Wagner2,
Mathias Winterhalter2, Haifeng Yuan10 and Giovanni Zifarelli32
1

Nano Life Science Institute (WPI-NanoLSI), Kanazawa University, Kanazawa, Japan
Department of Life Sciences & Chemistry, Jacobs University, Bremen, Germany
3
Ionovation GmbH, Osnabrück, Germany
4
MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine, University of Oxford,
Headley Way, OX3 9DS Oxford, United Kingdom
5Francis Crick Institute, London, United Kingdom
6
INM—Leibniz Institute for New Materials, 66123 Saarbrücken, Germany
7
Saarland University, 66123 Saarbrücken, Germany
8
Dpto. Física de la Materia Condensada Universidad Autónoma de Madrid 28049, Madrid, Spain
9
Instituto de Física de la Materia Condensada IFIMAC, Universidad Autónoma de Madrid 28049,
Madrid, Spain
10
KU Leuven, Department of Chemistry, B-3001 Heverlee, Belgium
11
Institute of Applied Optics, Friedrich-Schiller University, Jena, Germany
12
Leibniz Institute of Photonic Technology (IPHT), Jena, Germany
13
Department of Mechanical Engineering, University of Wisconsin-Madison, 1513 University Ave,
Madison, WI 53706, United States of America
14
Kennedy Institute for Rheumatology, University of Oxford, Oxford, United Kingdom
15
Debye Institute, Utrecht University, Utrecht, Netherlands
16
Department of Cell Biology, University of Groningen, University Medical Center Groningen,
Groningen, Netherlands
17
Division of Structural Biology, Wellcome Trust Centre for Human Genetics, University of Oxford,
Oxford, United Kingdom
18
Centre of Structural Systems Biology Hamburg and University of Hamburg, Hamburg, Germany
19
Heinrich-Pette-Institute, Leibniz Institute of Virology, Hamburg, Germany
20
Imaging Physics, Delft University of Technology, Delft, Netherlands
21
Department of Biochemistry, University of Oxford, Oxford, United Kingdom
22
Section Cell Biology, Center for Molecular Medicine, University Medical Center Utrecht, Utrecht
University, Heidelberglaan 100, 3584CX Utrecht, Netherlands
23
Centre for Advanced Imaging, The University of Queensland, Brisbane, QLD 4072, Australia
2

33

Authors to whom any correspondence should be addressed.

Original content from this work may be used under the termsof the Creative Commons Attribution 3.0 licence. Any further
distribution of this work must maintain attribution to the author(s) and the titleof the work, journal citation and DOI.
1361-6463/18/443001+42$33.00

1© 2018 IOP Publishing Ltd

Printed in the UK

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001
24

University Hospital Jena, Jena, Germany
Faculty of Medicine, Saarland University, 66421 Homburg, Germany
26
University of Göttingen, Third Institute of Physics—Biophysics, 37077 Göttingen, Germany
27
KU Leuven, Department of Bioscience Engineering, B-3001 Heverlee, Belgium
28
University of Göttingen, Institute for X-Ray Physics, 37077 Göttingen, Germany
29
SmarAct GmbH, Schütte-Lanz-Str. 9, D-26135 Oldenburg, Germany
30
Institute for Theoretical Physics and BioQuant, Heidelberg University, Heidelberg, Germany
31
School of Biochemistry, University of Bristol, Bristol, United Kingdom
32
Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
25

E-mail: christian.eggeling@rdm.ox.ac.uk, J.P.Hoogenboom@tudelft.nl and Schaap@smaract.com
Received 9 February 2018, revised 14 June 2018
Accepted for publication 1 July 2018
Published 31 August 2018
Abstract

Developments in microscopy have been instrumental to progress in the life sciences, and many
new techniques have been introduced and led to new discoveries throughout the last century. A
wide and diverse range of methodologies is now available, including electron microscopy, atomic
force microscopy, magnetic resonance imaging, small-angle x-ray scattering and multiple superresolution fluorescence techniques, and each of these methods provides valuable read-outs to meetthe demands set by the samples under study. Yet, the investigation of cell development requiresa multi-parametric approach to address both the structure and spatio-temporal organizationof organelles, and also the transduction of chemical signals and forces involved in cell–cellinteractions. Although the microscopy technologies for observing each of these characteristics
are well developed, none of them can offer read-out of all characteristics simultaneously, which
limits the information content of a measurement. For example, while electron microscopy isable to disclose the structural layout of cells and the macromolecular arrangement of proteins,it cannot directly follow dynamics in living cells. The latter can be achieved with fluorescencemicroscopy which, however, requires labelling and lacks spatial resolution. A remedy is tocombine and correlate different readouts from the same specimen, which opens new avenues tounderstand structure–function relations in biomedical research. At the same time, such correlative
approaches pose new challenges concerning sample preparation, instrument stability, regionof interest retrieval, and data analysis. Because the field of correlative microscopy is relatively
young, the capabilities of the various approaches have yet to be fully explored, and uncertainties
remain when considering the best choice of strategy and workflow for the correlative experiment.
With this in mind, the Journal of Physics D: Applied Physics presents a special roadmap on
the correlative microscopy techniques, giving a comprehensive overview from various leading
scientists in this field, via a collection of multiple short viewpoints.
Keywords: correlative microscopy, fluorescence microscopy, x-ray microscopy, electron
microscopy, magnetic resonance imaging, atomic force microscopy, super-resolution
microscopy(Some figures may appear in colour only in the online journal)

Contents
Integrated light and electron microscopy
Multiscale multimodal multicolor microscopy
Super-resolution CLEM: from room temperature to cryo-imaging
CLEM probes
CLEM in chemistry and catalysis
Fluorescence and electron microscopy of membrane proteins within intact cells in liquid
Correlative light and volume electron microscopy

2

4
7
9
11
13
15
17

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Force spectroscopy and single molecule fluorescence microscopy: unpacking single viruses
High-speed atomic force microscopy and light microscopy
Traction force microscopy
Super-resolved traction force microscopyElectrophysiology on lipid bilayers combined with fluorescence imaging and spectroscopyVoltage-clamp and patch-clamp fluorometry: studying ion channels and transporters with light
Fluorescence and magnetic resonance imaging
Fluorescence microscopy and scanning small-angle x-ray scattering: imaging of biological cells
Acknowledgments
References
List of abbreviations

20
22
25
28
30
32
34
36
38
38

QD
Quantum dot
NSOM
Near-field scanning optical microscopy
PFS
Point spread function
FCS
Fluorescence correlation spectroscopy
PMT
Photomultiplier
MRI
Magnetic resonance imaging
fMRI
Functional magnetic resonance imaging
dfMRI	Diffusion functional magnetic resonanceimaging
NMR
Nuclear magnetic resonance
SAXS
Small angle x-ray scattering
ROI
Region of interest
2D
two dimensional
3D
three dimensional
PIV
Particle image velocimetry
SPT
Single particle tracking
PCA
Principal component analysis
TFM
Traction force microscopy
STFM	
Super
resolution
tractionforce
microscopy
HLB
Horizontally oriented bilayer
OSTR
Optical single transporter recording
VCF
Voltage-clamp fluorometry
PCF
Patch-clamp fluorometry
BOLD
Blood oxygenation-dependent

CLEM
Correlative light and electron microscopy
TEM
Transmission electron microscope
SEM
Scanning electron microscope
FIB
Focussed ion beam
CL
Cathodoluminescence
EDX
Energy-dispersive-x-ray analysis
EELS
Electron energy loss spectroscopy
nano-SIMS	Secondary ion mass spectroscopy at thenanoscale
STEM
Scanning transmission electron microscopy
ESEM
Environmental scanning electron microscopy
EMPIAR
Electron microscopy public image archive
LM
Light microscopy
FRET
Förster resonance energy transfer
STORM	Stochastic
opticalreconstruction
microscopy
PALM
Photoactivated localization microscopy
AFM
Atomic force microscopy
FM
Fluorescence microscopy
GFP
Green fluorescent protein
FP
Fluorescent protein
NA
Numerical aperture
SiN
Silicon nitride3

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Integrated light and electron microscopy
Hans C Gerritsen1 and Jacob P Hoogenboom2
1

Debye Institute, Utrecht University, Utrecht, Netherlands
Imaging Physics, Delft University of Technology, Delft,
Netherlands
2

Status. The integration of a light microscope (LM) and anelectron microscope (EM) into a single apparatus has been
pursued for doing correlative light and electron microscopy (CLEM) experiments since the 1980s. Technologicaladvances and the renewed interest for CLEM have led toseveral novel, improved systems, both for transmission EM(TEM) and scanning EM (SEM) (see overviews [1, 2]). Integrated CLEM approaches improve correlation or image registration accuracy, facilitate the retrieval of (rare) regions ofinterest, reduce CLEM operation times, and/or avoid sample
contamination in (cryo-) transfer [3]. Potential drawbacks for
integrated microscopy are the need for fluorescence preservation during preparation for EM and in vacuum, and autofluorescence of some resin materials. Preparation schemes
using reduced concentrations of osmium and metal salts [4–6]
as well as osmium-resistant genetic labels [7] have evolved,
while extension to integrated cryo-microscopy [8, 9] may alleviate all these issues.
Several types of integrated microscopes are now commercially available and increasingly finding their way into
the laboratory environment. These can be roughly divided in
two variants: systems where EM and integrated LM share thesame field of view (figures 1(a) and (c)), and those where thesample needs to be translated or rotated within the vacuum
chamber in between imaging with both modalities (figures
1(b) and (d) [2]. In the first variant, image registration does
not need fiducial markers (see figure 2), but can be doneusing so-called cathodoluminescence pointers, which can be
extremely accurate (<10 nm) and automated [10], but sample thickness is limited by the depth of view of the LM. The
latter case is applicable to (cryo-)TEM and can be used in
SEMs, removing the restriction on sample thickness. Eithercase may hold considerable benefits to target key challenges
in EM and CLEM. Larger samples are used in volume-EM
where the sample is trimmed with focused ion beams (FIB) or
in situ microtomes [11, 12]. Automated and highly accurate
integrated CLEM may be key for superresolution (SR) fluorescence localization of bio-molecules in EM images [13], forlocating and trimming sections for sub-nm resolution structural cryo-EM [12], and for large-scale serial section EM [11].
A recent demonstration of integrated SR fluorescence CLEM
showed a localization accuracy of 50 nm [14], comparable toroutine stand-alone SR experiments.

Figure 1. Schematic indication of realizations for integrated LM
inside ((a), (b)) scanning or ((c), (d)) transmission EMs. Designs
can be distinguished based on whether ((a), (c)) both microscopes
share the same field of view, or (b) a translation, or (d) rotation isneeded, to switch from light to electron microscopy and vice versa.
Electron beam is indicated in green, light beam in blue.

EM. Ultimately, localization accuracy should be comparable
to the nanometer resolution of EM, effectively adding fluorescence contrast to EM. Current localization based SR techniques, such as photoactivated localization microscopy andstochastic optical reconstruction microscopy, routinely obtain
resolutions of 20–50 nm. However, below 10 nm resolution,
SR-EM registration accuracy and/or distortions induced bysample preparation become dominant. At this length scale,integrated microscopes and optimized ‘integrated’ specimen
preparations are likely to yield the best results, because dist­ortions due to specimen handling can be avoided and registration accuracy is high.
Cryo-CLEM. Revolutionary developments in cryo-EM andelectron tomography (ET) resulted in near-atomic resolution that enables resolving the internal structure of proteins.
A major advantage of EM over crystallography approaches
would be imaging the structure of a protein in its native, cryofixed environment. The holy grail in cryo-EM is to pinpointa protein of interest in a cryo-fixed specimen and cut out asufficiently thin slice (100–200 nm) containing this protein fortransfer to cryo-EM/ET. FIB SEMs are the tool of choice forslicing, and cryo-fluorescence microscopy can highlight theprotein of interest. A major challenge is to reach the precisionneeded for targeted 100–200 nm slicing in cryo-fixed cells,
which can most probably only be reached with cryo-integrated
fluorescence FIB-SEM. Challenges include accurate 3D correlation, especially considering the poor depth resolution in(confocal) microscopes and optical distortions.

Current and future challenges.
SR CLEM.
Combining SR fluorescence with EM holds

the promise of precisely pinpointing molecules that cannot
be labelled for EM in EM images. SR CLEM opens the door
to functional imaging of, for instance, specific lipids, ions or
enzymatic activity in the ultrastructural image obtained with

Volume- and high-throughput EM. The throughput in EM

has evolved considerably. Large areas can be covered thanks
to technical developments, e.g. increasing the size of sections cut by the microtome and using SEM for seamless4Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

applications of fluorescence preserving EM sample prep­
aration, on-section immuno-labelling, and reduction of resin
auto-fluorescence. For block-face approaches, fiducial markers or calibration structures for 3D registration need further
development.
Advances in science and technology to meet challenges.
Fluorescence and photo-switching under EM conditions.

Optimized ‘integrated’ sample preparation is a common challenge to all of the approaches detailed above. For SR fluorescence, three hurdles need further attention. First, fluorescencehas to survive fixation and other EM preparation steps, which
has been achieved [1, 4–7], but needs a wider palette. Strong
fixation and staining in 3D block-face requires the development of milder fixation procedures compatible with CLEM.
Second, fluorescence and photo-switching has to be preservedin vacuum (dehydrated state); for most genetic fluorophores
this results in low fluorescence quantum yield. Variable-pres­sure EM provides an alternative, but is time-consuming [14].
Techniques relying on direct electron beam excitation (cathodoluminescence) may provide alternatives (see section 2), but so
far lack sufficiently small (<20 nm), stable probes. In addition,
optimized genetic photo-switchable probes may provide a solution. Third, background fluorescence from resin may become
problematic at low fluorescence signals (SR) or thick samples(3D CLEM). This requires developments of low auto-fluorescence resins and use of long wavelength fluorescent probes.
Much of these issues are alleviated with cryo-fixation,
but photo-switching may be difficult, and illumination powers needed for SR microscopy may lead to local melting.
Again, optimized fluorescent proteins for cryo-conditions are
required, combined with the development of novel, cryo-specific SR techniques.

Figure 2. Examples of (a)–(c) fiducial and (d)–(f) non-fiducial
based image registration in integrated microscopes. (a) FM image
in TEM (implementation according to figure 1(d)) of Tokuyasu
sections of HeLa cells transfected with LAMP-1-GFP. Nuclei areshown in blue (DAPI), LAMP-1-GFP in green and fiducials in red.
(b) Overlay of ROI (boxed area in (a)) of fluorescence and TEM
images. (c) Zoom in on LAMP-1-GFP rich area. Fiducials consistof silica particles with a 15 nm gold core and a 40 nm fluorescently
labeled silica shell. Overlay accuracy is about 30 nm. (d) FM image
in SEM (implementation according to figure 1(a)) of rat pancreas
sections, immuno-labelled after embedding in epon to show nuclei
in blue (Hoechst), guanine quadruplexes in light blue (Alexa488),
and insulin in orange (Alexa594). (e) SEM image of the ROI (boxedarea in (d)). (f) Overlay of fluorescence from the ROI with the SEM
image. The overlay (<20 nm accuracy) is obtained via an automatedregistration procedure between both microscopes [10]. Scale bars
are 10 µm in (a), (d), 2 µm in (b), (e) and (f), and 0.5 µm in (c).

Registration and depth resolution in 3D. While integrated

microscopes allow for automatic image registration for thin
2D samples, registration of thicker 3D samples (volume
CLEM) poses challenges. Novel fiducials are needed, which
are bright enough in sample blocks, heavily stained with
osmium and other metals and yield backscattered electron
visibility in the stained blocks. Also, techniques to accountfor optical distortions (due to refractive index mismatch, forexample) such as adaptive optics, need to be incorporated.
Fluorescence localization in 3D requires optical sectioning, e.g. using an integrated confocal microscope. A major
challenge is to achieve depth resolution matching that of volume-EM techniques, i.e.  ⩽100 nm.

imaging, as well as developments in software and automation,such as automated image acquisition and stitching. Volumes
can be covered by an automated collection of thin serial sections and these imaging sections sequentially, or by imaging
the upper face of the resin block followed by in situ trimming
using an integrated microtome or FIB-SEM (see section 7).
The recent acquisition of a zebrafish brain using serial-section
SEM constitutes a hallmark example of what can be achieved
with volume-EM [15]. However, data acquisition took over
200 full days of SEM operation, highlighting the need to pinpoint regions of interest to cut redundancy in acquisition, forwhich integrated CLEM seems excellently suited. Paired withthe high-accuracy fluorescence-to-EM registration that can beobtained consistently over large areas, integrated microscopes
seem particularly suited to improve throughput and functional
mapping in serial sections volume-EM. Instrumentation
seems to be in place, but automation, especially in fluorescence recognition and unattended acquisition, needs development. Challenges also remain in further, more wide-spread

Automation. For 2D and 3D serial sections, instrumentation
seems to be in place to move to a high throughput acquisitionof large, accurately overlaid CLEM datasets [16]. Effort isrequired in the automation of data acquisition, drift and focus
corrections and section recognition, all geared towards unattended, 24/7 operation. Further along the way, the automatedrecognition of target areas for EM, based on in-section fluorescence expression, may also help to enhance throughput inserial-section CLEM.

5

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

microscopes may allow recording of precisely overlaid datasets of functional fluorescence and structural electron datacrossing scales from the multi-cellular down to the molecularlevel.

Concluding remarks

A variety of integrated microscopes have emerged in recent
years. Key application areas for integrated microscopy are in
CLEM with super resolution fluorescence, CLEM for cryo- orvolume-EM, and high-throughput CLEM based on serial sections. Sample preparation protocols and fluorescence probes
for integrated CLEM are available, but in general, solutions
require both further and broader optimization of genetic andorganic fluorophores towards EM conditions (cryo, vacuum,use of osmium and other stains), methods for 3D registration and resolution matching, and development of automateddata acquisition strategies. With these steps made, integrated

Acknowledgments
We thank our group members and collaborators for input
and discussions. We acknowledge funding from Microscopy
Valley, a research program supported by NWO-TTW
Perspectief voor de Topsectoren (projects 12713, 12714 and
12715). J P H has a financial interest in Delmic BV, a company producing integrated microscopes.

6

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Multiscale multimodal multicolor microscopy(1)	Large ROIs—data avalanche: many initiatives try to handle
and publish large imaging datasets. Our lab has pioneered
placing large-scale EM maps of a variety of cells, tissues andmodel organisms at full resolution with open access at www.
nanotomy.org: the large-scale data at the full resolution of
figure 3 are available on this website. Similar initiatives
are being undertaken by others and are referred to on thiswebsite. Current developments in data sharing, analysis, andinterpretation—like automated recognition and machinelearning software—will evolve in the next decade and solve
data handling challenges. Increasingly, open access to highcontent multidimensional microscopic data will pave thepath for multimodal analysis of the high content datasets.
Metadata definition and (online) representation of such
multi-dimensional and multi-parameter datasets needs tofurther develop to meet international consensus.
(2)	Localization precision of many molecules and structure:with the recent broad implementation of scanning EM inlife sciences, going beyond surface characterization withsecondary electron detection, but also using backscatter
detectors, transmission electron detection and even fluorescence in hybrid LM/EM microscopes [1] (section 1),
new opportunities to analyse tissues arise. Using the electron beam to generate signals in a scanning EM allows
us to achieve lateral nanometer range localization, even
if the detected signals are photons, for instance (table 1).
Such as in light microscopy, axial resolution matching
the lateral resolution will remain a challenge. Correlative
microscopy will develop towards using more probes [19]and detectors to define the localization of molecules using
endogenous signals or new probes, but not depending on
fluorescence per se.

Ben N G Giepmans

Department of Cell Biology, University of Groningen, University
Medical Center Groningen, Groningen, Netherlands
Status. Correlative light microscopy and electron m­ icroscopy(CLEM) is a key approach to studying structure–function relationships in cell biology. CLEM allows a biological process
and building block (molecule, organelle, cell) to be identified
and dynamically studied using fluorescent markers, followedby high-resolution analysis of the ultrastructural context with
EM. In the past few decades, sample preparation steps, technical approaches, probes, microscopes and image analysis havebeen optimized to make CLEM a routine approach applied bymany labs to date [1].
The strength of biomedical EM obviously lies in revealing
the ultrastructure and detecting targets at a biomolecular scale,
which is typically a few nanometers (nm) for proteins. In contrast, in fluorescence microscopy only the targets (typically
restricted to three) are highlighted, with limited informationon the non-labelled structures. The resolution and localization
precision of fluorescence imaging has long been restricted to a
sub-micrometer scale because of the diffraction limit. The riseof nanoscopy, i.e. light microscopy beyond the diffraction limit,
directly showed a high potential for defining the localization oforganelles with high precision (<0.1 µm; see [1, 17]). Recently,
Hell et al [17] developed ‘MINFLUX’, leading to nm-precision
localization of labelled fluorescent molecules with a lateral resolution of 6 nm, enabling cell biology at the ultimate scale of biomolecules, but only for one or two targets at a time.
Current and future challenges. Although all microscopic pre-Advances in science and technology to meet challenges. The

embedding labelling procedures often lead to the extraction of
biomolecules, only EM painfully shows the effect on the ultrastructure [18]. Well-established ways to circumvent cellular
damag, by introducing probes are by using either geneticallyencoded tags and/or small molecules that do not need permeabilization [1] or post-embedding labelling, e.g. figure 3. The
concession in CLEM for sample preparation, not optimal for
either modality, also counts for the light microscopic analysis.
Typically, fluorescence is poorly retained during EM sample
preparation focussed on retaining optimal ultrastructure,
although probes and procedures are being developed to retain
fluorescence during embedding (reviewed in [1]), or fluorescent labels can be targeted post-embedding.
Another challenge in CLEM is to deal with large datasets. The successes of CLEM approaches have resulted in ademand to image large regions of interest (ROIs), typicallyeasily performed with LM, at the typical resolution for EM(nm-scale). In dynamic vital microscopy followed by EM, thisgap will not be easy to routinely solve, especially because of
the difficulty of correlating data in the axial direction. In fixedsamples, matching the scales between the modalities demands
large-scale EM [1], which leads to an avalanche of data andthe current quest is to identify or fingerprint biomolecules in
the nm-range.

endeavour the field made, ranging from sample preparation tomultimodal microscopes with a variety of detectors, will lead to‘correlative microscopy’ that will increasingly use the resolution that can be achieved by the electron beam, using analysis
that leads to unique identifiers of molecules in EM-imaging.
Cathodoluminescence (CL). In the multimodal microscopes,

when electrons hit CL molecules, photons can be collected
resulting in localization defined by the electron beam [20]. In
addition, new analytical methods are being pioneered, which
are based on using an electron beam or ion beam to achieve
the required lateral localization precision.
Energy dispersive x-ray analysis (EDX). Elemental fingerprinting using EDX was described in a tour-de-force way by
Somlyo et al, 40 years ago [21], who revealed subcellular distribution of elements in muscle using spectroscopic methods.
Development in EDX detectors and computer software nowadays allows (semi)routine EDX imaging of 1k  ×  1k pixel
areas, to fingerprint biomolecules and probes in the context of
ultrastructure (figure 3) [22].
Electron energy loss spectroscopy (EELS). In parallel,elemental analysis is performed using EELS. Leapman et al7Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 3. ‘ColorEM’ using elemental analysis by energy dispersive x-ray imaging. ColorEM: label-free (P), paint (Os) and labeling DNA(Au) and peptides (Cd) is compatible. (a) Part of an islet of Langerhans immuno-labeled for structures in DNA (10 nm gold) and insulin(QD). (b) Overlay image of Au (red), Cd (green), Os (yellow) and P (blue) allows identification of G4 structures (gold labels) and insulin(Cd). Note the localization of Au to heterochromatin enriched in P, whereas the Cd signal is enclosed within a combination of Os rings andP that likely identifies phospholipid membranes of the vesicles. Large scale data and full resolution data is available via www.nanotomy.
org; Reproduced from [22]. CC BY 4.0.
Table 1. Approaches for CLEM and ColorEM.

FLM
EM
CL
EDX
EELS
nanoSIMSaIn

Detected

Oppertunity

Limitation

Photons
Electrons
Electrons
Electrons
Electrons
Ions

Visible photons
Electrons
Visible photons
X-raysElectrons‘Molecules’

Live cells, large area
Ultrastructure, unbiased
HIGH resolution
Endogenous and probes
Endogenous and probes
Endogenous andprobes

Needs probes, resolutiona
Limited probes, grey scale
Needs development
Undefined molecule
Undefined molecule
Resolution

Except for nanoscopy; see text for details.

pioneered EELS imaging in tissue to detect endogenous elements [23]. Similarly, EELS TEM allows the detection of particles enriched in certain elements targeted for labelling, suchas quantum dots [24]. Recently, Tsien et al used lanthanidesenriched molecules that can be deposited using specificprobes to perform two-color EELS, to discriminate targetedmolecules and biostructures of interest [25].

between nanoscopic fluorescence microscopy and EM is now
feasible. CLEM will develop to make the workflow more convenient, faster and more generically available. The data sets will belarger, and protocols for reuse of data will be developed as exemplified by several initiatives to already share large-scale EM data.
The major breakthrough in correlative microscopy in thedecade to come is predicted to be the generic use of multimodal
microscopic imaging. Microscopists will make better use of
the electron beam or ion beam, to generate signals that fingerprint or identify biomolecules and structures, either directly
or indirectly, using to-be-developed probes and bypassing thediffraction limit of light microscopy. These developments willlead to multidimensional EM with a pleiotropy of signals andmolecules detected at nm-scale precision and reveal many
current secrets underlying the regulation of life.

Secondary ion mass spectroscopy at the nanoscale (nanoSIMS). Elemental analysis using the electron beam leads tofingerprinting of which class of biomolecules are present. However, this does not typically identify molecules, like defining
which protein is present. Using an ion beam instead allows
the performance of nano-SIMS [26]. Thus, a map of isotopelabeled proteins can be identified at up to 50 nm resolution. The
electron-beam and ion-beam techniques hold great promise to
become standard tools in correlative microscopic imaging, as
they allow imaging of characteristics that lead to molecular
identification of biomolecules in an unbiased manner.

Acknowledgments
I thank my team members and Jacob Hoogenboom, Delft
University of Technology, for discussions. Our work relevantto this paper is supported by the Netherlands organization for
scientific research (ZonMW 91111.006; STW Microscopy
Valley 12718; TTW15315).

Concluding remarks. CLEM has developed to a semi-routine
technique. Major bottlenecks, like retaining fluorescence in EMprepared samples have been overcome. Thus, a combination
8

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Super-resolution CLEM: from room temperature
to cryo-imaging
Rainer Kaufmann1,2,3 and Kay Grünewald1,3,4
1

Division of Structural Biology, Wellcome Trust Centre for
Human Genetics, University of Oxford, Oxford, United Kingdom
2
Department of Biochemistry, University of Oxford, Oxford,
United Kingdom
3
Centre of Structural Systems Biology Hamburg and Universityof Hamburg, Hamburg, Germany
4
Heinrich-Pette-Institute, Leibniz Institute of Virology,
Hamburg, Germany
Status. Super-resolution

correlative light and electron
microscopy (super-resolution CLEM) is a quickly evolving
addition to the CLEM field that presents a true game-changer(see figure 4). Before, the large resolution gap between conventional fluorescence microscopy (FM) and EM did typically only allow for FM-based rough localization of areas or
events of interest to be subsequently targeted by EM imaging.
Despite the complementarity of both microscopy techniques,true correlative imaging was not possible before the introduction of super-resolution FM methods.
With the growing number of super-resolution methods andthe plethora of EM protocols, the complexity and choices ofcombining both imaging modalities has tremendously increased
over the past few years, since the first demonstration of superresolution CLEM by Betzig et al in 2006 [27]. Microscopy
hardware has significantly advanced for both imaging modalities, and the challenges in super-resolution CLEM, as Shtengel
and Hess have recently pointed out [28], are mainly in sample
preparation. Often, the typical protocols for EM are incompatible with super-resolution FM requirements and vice versa.
Recently, various workflows addressing these limitations havebeen developed, enabling there now to be more possibilities
for combinations of super-resolution CLEM. For example, thedevelopment of an OsO4 resistant photoactivatable fluorescentprotein (FP) allowed the introduction of super-resolution FM
compatible markers into the typical EM fixation workflow,which, in its standard protocol, destroys the fluorescence [7].
In an alternative approach, the freeze substitution and resin
embedding procedure was adapted to maintain the fluorophore’s photo-switching capabilities, enabling in-resin superresolution CLEM using standard FPs [29].
For cryo-conditions, the challenges, and hence the current
status of super-resolution CLEM, are very different. While aresolution down to the Ångstrom range for biological samples
is possible on the cryo-EM side, the resolution of cryo-FM
is severely limited by respective technical requirements [30].
Most importantly, the lack of high-numerical aperture (NA)cryo-immersion objective lenses reduces the resolution toabout half of what is achievable in conventional FM at ambient temperatures. While the feasibility of super-resolution
cryo-CLEM has conceptually been demonstrated [31, 32],
huge technical and photo-physical challenges [33] are cur­rently hindering routine biological applications of this potentially very powerful technique.

Figure 4. Overlay of fluorescence signal of mVenus-labelled
EphA2 protein imaged by conventional FM (top) and superresolution single molecule localization microscopy (SMLM)(bottom) with a TEM image in a freeze substituted and resin
embedded HEK293T cell using a dedicated super-resolution
CLEM protocol [34]. The structural resolution of in-resin SMLM
was approx. 50 nm with an average single molecule localization
accuracy of 17 nm. Reproduced from [29]. CC BY 4.0.

Current and future challenges. At ambient temperatures,
super-resolution CLEM requires balancing requirements ofresolution in FM, contrast in EM and structural preservation[7, 28, 29, 33, 35]. For example, Johnson et al [29] reported
that tannic acid, the key component for preserving the photoswitching of FPs in their protocol, had an antagonistic effecton the achievable FM resolution and structural preservation of
the sample. Further, due to structural changes during the EM
fixation, dehydration, embedding and even imaging processing— which are typically performed after FM data acquisition—the correlation of FM and EM images can suffer a fairly
large uncertainty regarding the relative positions of fluorescent labels and EM structural features [28]. Other approaches,such as the above-mentioned OsO4-resistant photo-activatable FP [7] or preservation of photo-switching of FPs duringfreeze substitution and resin embedding [29], minimize thestructural changes between super-resolution FM and EM dataacquisition, by allowing for imaging on specimens after EM
preparation. This results in improved correlation accuracy, but
imposes limits regarding the choice of suitable fluorescentmarkers.
Super-resolution cryo-CLEM, on the other hand, is cur­rently still much more limited by technical challenges [30].
Cryo-FM suffers from the fact that, so far, no high-NA immersion objective lens has been developed for cryo-conditions.
This restricts not only the optical resolution, but also the9Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Optimization of these fluorophores and dedicated protocols
will, moreover, help to increase the resolution of FM. Parallel to this, it is also important to minimize structural changes,
both those that might occur between FM and EM imaging,and those which might become problematic in general for the
biological interpretation of the data. If the correlation acc­
uracy becomes the limiting factor in super-resolution CLEM,
there is no point in pushing the FM resolution even further,and evidently, both imaging modalities will be impaired if
structural artefacts arise.
Super-resolution CLEM under cryo-conditions currently
has many more areas that require improvements. The crucial parameter of SMLM super-resolution imaging is typically the photo-switching of the fluorescent markers. This isvery poorly studied and understood under cryo-conditions,and is probably the factor that would have the biggest impact
on the quality and usability of super-resolution cryo-CLEM.
Another (often neglected) challenge is the problem of devitrification. This has also currently been the reason why stimulated-emission-depletion, requiring at least in its standard
form very large laser powers, has not yet been successfully
applied in cryo-CLEM applications [30]. A generally applicable solution is required, that overcomes the current limitation to subset samples, and is fully compatible with cryo-EM
imaging. Super-resolution cryo-CLEM loses its justification
if the advantages of cryo-EM—structural preservation andhighest resolution—cannot be maintained. Encouragingly,
several groups are now actively tackling these challenges anda noticeably wider community of hardware providers havebecome interested.

Figure 5. Top: cryo-SMLM of a mitochondrial protein labelledwith clover in a whole vitrified COS7 cell before thinning byfocused ion beam milling (for method see [12]). For cryo-SMLM alaser intensity of approx. 1.6 kW cm−2 was used. Bottom: cryo-EM
images after thinning of the areas (blue rectangles) indicated in the
cryo-SMLM image. Segregation artefacts and Bragg reflections
indicate devitrification (ice crystal formation) caused by cryoSMLM imaging.

Concluding remarks. Super-resolution CLEM is becom-

detection efficiency, that is critical for super-resolution methods. Another aspect crucial for single molecule localization microscopy (SMLM) based super-resolution FM is theability of photo-switching of fluorescent molecules, but theunderlying photo-physics is only poorly understood for cryoconditions. So far, only SMLM methods have been used forsuper-resolution cryo-CLEM [31, 32]. These require a certain level of laser intensity for switching the fluorescent molecules to achieve super-resolution. Accordingly, the biggest
challenge currently preventing the wider biological application is sample devitrification by local warming, resulting in
the transition of amorphous ice to a crystalline form, thereby
destroying the biological structures [30–32] (figure 5). Hence,so far, successful super-resolution cryo-CLEM has only beenachieved by using cryo-protectants and/or formvar coatedgrids [31, 32], which are both not ideal for cryo-EM [30].

ing an established method, for which current developments
are focusing on making it compatible with a broader rangeof samples, and overcoming the limitations imposed by the
choice of fluorescent markers or other constraints during sample preparation. Currently, super-resolution CLEM methods
based on freeze substitution and resin embedding provide, formost biological applications, the best compromise of resolution on the FM side, and structural preservation on the EM
side. In contrast, super-resolution cryo-CLEM is a techniquethat is just emerging and still requires significant improvements to turn it into the powerful biological tool it promises tobe. If proven practically feasible for a wider specimen range,
it will surely become the method of choice, due to the super­
ior structural preservation of vitrified samples and resolution
on the EM side, and might even take the full advantage ofimproved fluorophore properties under cryo-conditions.

Advances in science and technology to meet challenges. For
the advancement of super-resolution CLEM under ambientconditions, improvement of the sample preparation protocols
remains the most important aspect. The development of more
fluorescent markers or protocols to preserve their capabilities for super-resolution imaging will make super-resolution
CLEM suitable for a wider range of biological questions.

Acknowledgments
We acknowledge support from the Wellcome Trust(107806/Z/15/Z to KG and 107457/Z/15/Z to Micron
Oxford), HFSP (RGP 0055/2015 to KG) and CRUK (A17721
to E Yvonne Jones).

10Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

CLEM probes(2)	Single and dual modality exogenous probes:(a)	Single fluorescent or gold probes that are photoconverted or can be visualised with multiple microscope
techniques. These include the recent development oflabelling non-protein molecules via click chemistry[36, 37].
(b)	Probes with a metal core and inherent fluorescent
properties—lanthanides/quantum dots [38]. These
probes are very well suited for CLEM experiments
as they are inherently fluorescent and electron dense.
But due to their relatively large size, penetration andfunctionalisation can be significant challenges.
(c)	Probes with a metal core conjugated to a fluorescentmoiety—fluoronanogold. Fluoronanogold with its
1.4–1.8 nm gold particle conjugated next to a fluorescent molecule is much smaller than a quantum dot
and therefore penetrates better but is not directly visible and requires silver or gold-enhancement.

Paul Verkade1 and Lucy Collinson2
1

School of Biochemistry, University of Bristol, Bristol, United
Kingdom2
Francis Crick Institute, London, United Kingdom
Status. In almost all imaging techniques, the selection of theright probe(s) is an essential part of the workflow. Probes areused to mark the region of interest and to reveal functional
information about the biological processes under investigation.
For correlative imaging, where different imaging modalities
are combined in one experiment, it is possible to use a probethat is visible in one modality (e.g. fluorescence microscopy)
and to track the physical location of the probe, and analyse
the properties of the sample in another imaging modality (e.g.
electron microscopy). However, if the probe is not visible in thesecond imaging modality, there will be a degree of uncertainty
in the correlation. A true correlative probe must be visible ineach imaging modality used. This is a challenge because light,
electron, x-ray and force microscopes have fundamentally different contrast mechanisms and sample interactions.
The ideal probe will reach its target on, or in, cells andtissues with no cytoxicity and no adverse effects on ultrastructure or on the biological process under study. It will havemaximal contrast in all imaging modalities, and will be smallenough to localise the macromolecule to a cellular structurewith an accuracy equal to the size of that molecule.
Though there is a wealth of knowledge on probes and there
is continuous development of new probes, the ideal universal
probe is yet to be designed (figure 6) [35]. Realistically, it isunlikely that a single probe will be suitable for every different correlative imaging workflow, and researchers will need toevaluate which probe is best suited for each scientific question.
There are, however, great opportunities in the field for developments, both in the probes themselves and new technologies, todetect existing probes across the different imaging modalities.

3.	Dual modality genetic probes:(a)	Probes that can be converted from a fluorescent to anelectron dense signal using photo- or chemical conversion—mini singlet oxygen generator (miniSOG).
Probably superseded by APEX, mini-SOG can still be
used as a CLEM probe as it is fluorescent (although
not very bright) and can generate a DAB precipitate
because of its singlet oxygen generating properties.
(b)	Probes with two genetic tags added together, one
fluorescent and one convertible to an electron dense
product—horse radish peroxidase (HRP) or APEX
with a fluorescent protein (e.g. GFP) [39]. APEX is amuch better SOG than mini-SOG and hence a combination with a better fluorescent molecule than mini-SOG
provides a very useful combination as CLEM probes.
(c)	Probes with two genetic tags added together, one
fluorescent and one metal-binding protein—e.g.
metallothionein or ferritin with a fluorescent protein(e.g. GFP). To potentially improve the localisation
precision of the EM marker from a precipitate to aparticulate marker, APEX can be replaced by a metalbinding protein such as ferritin.

Current and future challenges. To date, most correlative lightand electron microscopy (CLEM) probes can be categorised
into three classes:(1)	Single modality genetic probes that are compatible withsample preparation for two or more imaging modalities:

Genetic probes have an advantage because no permeabilisation of the cell membrane is required to gain access tointernal structures, and so membrane integrity is preserved,
which is critical for structural studies by electron and x-ray
microscopy. However, expression levels must be carefully
controlled, as over-expression of fluorescently-tagged molecules often disrupts the structures and processes under study.
Smaller genetic probes are preferred since they are less likely
to cause misfolding or aggregation effects upon expression,
so a single mEos4/GFP/miniSOG construct may be preferredover a dual GFP-APEX of GFP-HRP construct.
Immuno probes avoid issues of probe expression, butvarying levels of membrane disruption, using detergents(triton) or toxins (saponin, digitonin), are required to allow
entry of the probes to the cell. The exception is in studies
of endocytosis or phagocytosis, where probes can be fed to(a)	Osmium-resistant probes for fluorescence and electron
microscopy—the fluorescent Eos derivative mEos4 hasrecently been described to withstand osmium treatment
during EM processing [7]. The mEos4 molecule is therefore still fluorescent in a well-stained resin-embedded
sample. In addition, it is compatible with super resolution photoactivated localization microscopy.
(b)	Fluorescent proteins—green fluorescent protein (GFP)
family [5]. These fluorophores survive a mild processing
regime that retains water and avoids osmium and epoxy
resins during embedding. By very precise alignment of
the images obtained via LM and EM using fluorescentbeads, the location of GFP tagged proteins can be determined within ~100 nanometers.
11

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 6. Example of a mismatch between fluorescence and gold coupling to proteins of interest. Epidermal growth factor coupled toAlexa488 and 10 nm gold particles internalised into HeLa cells shows bright fluorescence but only one gold particle at the region ofinterest. Transferrin coupled to Alexa488 and 5 nm gold particles, however, shows a lower fluorescence signal but numerous gold particles.
Quenching of the fluorophore and coupling efficiency may play a role in this mismatch. See also [14].

cells in targeted (via membrane receptors) or untargeted (via
fluid-phase uptake) feeding experiments. Care must also betaken here that binding to the receptor or fluid-phase uptake
does not adversely affect the trafficking pathways being
studied. Access of these probes becomes increasingly difficult as the sample size increases into the tissue and organism
domain. Other issues that must be considered include cytotoxicity, size and steric hindrance effects in dual labelling
experiments.

(2)	Advances in Molecular Biology—the use of knock-down
and re-expression, especially exploiting the Crispr/CAS9
system for the expression of genetically encoded probes
tagged to proteins of interest will be a major advance incontrolling off-target effects of those probes.
(3)	Advances in sample preparation—milder conditions
compatible with preservation of probe contrast between
imaging modalities [14].
(4)	Advances in microscope technology—new excitationand detection regimes compatible with existing and newprobes that enable ‘ideal probe’ conditions. Examples of
those are the direct detection of fluorescent probes in theelectron microscope using electron energy loss spectr­
oscopy or nanoscale secondary ion mass spectrometry,
Raman for probe-free element detection, or detection of
metal particles using photon detection techniques such as
cathodoluminescence, four wave mixing and interferometric cross-polarization microscopy [40].

Advances in science and technology to meet challenges. In
correlative imaging, similar to most (biological) research
fields nowadays, it requires the combined effort of scientists with multi-disciplinary backgrounds to best tackle thescientific problem. Only by combining our expertise in biology, chemistry, physics and microscopy will we be able togenerate better correlative imaging probes—a process that is
analogous to correlative imaging in leveraging the strength ofeach individual microscope, and combining them in a single
experiment to generate more than just the sum of each technique. Below, we highlight some of the specific areas that cancontribute to such development:

Concluding remarks. Most correlative imaging experiments
currently link two imaging modalities. There is great potential
in developing probes to link three or more imaging modalities[41], and in doing so, reach from clinical imaging (MRI, CT,
PET/SPECT, fluorescence image-guided surgery) through tothe molecular scale. The work must be focused around a suiteof important test-case biomedical research questions to ensurethat the probes are fit for purpose.

(1)	Advances in chemistry—targeted evolution of existing
genetic probes and design of new probes to resist the
sample preparation steps required to move between
imaging modalities [7]; further investigation of andimprovements in the production of homogenous nanoparticles of lanthanides; focused development of quantum
dots as imaging probes to improve electron density whilst
maintaining fluorescent intensity; development of small
molecules that can enter the cell without permeabilisationor cytotoxicity and that are visible in multiple imaging
modalities; investigation of new probe types with multiple contrast mechanisms (nanodiamonds, others).

Acknowledgments
This work was supported by the Francis Crick Institutewhich receives its core funding from Cancer Research UK(FC001999), the UK Medical Research Council (FC001999),
and the Wellcome Trust (FC001999).
12

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

CLEM in chemistry and catalysis
Elke Debroye1, Haifeng Yuan1, Johan Hofkens1,
Maarten B J Roeffaers2 and Kris P F Janssen1
1

KU Leuven, Molecular Imaging and Photonics, Departmentof Chemistry, Celestijnenlaan 200F, B-3001 Heverlee,
Belgium
2
KU Leuven, Centre for Surface Chemistry and Catalysis,
Department of Bioscience Engineering, Celestijnenlaan 200F,
B-3001 Heverlee, Belgium
Status. The nanoscale dynamics and complexity of hetero-geneous molecular systems carry over to larger length scales,
where phenomena such as diffusion, Brownian motion, crystal growth or charge carrier transport are all manifestations of
stochastic processes at the molecular level. While nanoscale
phenomena have a marked impact on the performance of catalytic systems [42], or the efficiency of novel energy harvestingmaterials [43], they are typically obscured by ensemble averaging and/or the limited time resolution of bulk characterization
methods. More advanced techniques to explore the intrinsic
nature of these materials at the nanoscale are therefore needed.
As an example, single-molecule and/or super-resolution fluorescence (SRF) imaging [42] allow chemical activity distributions to be studied at the single particle level, in turn providing
important information to understand the macroscopic performance of industrially applied heterogeneous catalysts [42, 44].
Although fluorescence microscopy affords spatial mapping ofchemical activity or the tracking of single molecules, often withnanometre precision and high time resolution, it is less suited toprovide the structural context that sets the stage for molecular
events and processes of interest (figure 7). It is for these reasonsthat correlative light and electron microscopy (CLEM), in its
various embodiments, has been successfully applied in biology
to reveal structure-activity relationships [1] and is now increasingly being applied in chemistry and catalysis research.
Using CLEM, reductive N-deoxygenation of resazurin by
NH2OH and oxidative N-deacetylation of amplex red by H2O2,
both yielding the highly fluorescent resorufin, were used asprobe reactions to map catalytic activity on gold nanoparticles
coated with mesoporous silica (Au@mSiO2) [45]. The sameprobe reaction was used to study the photocatalytic activity at
distinct surface facets of titanium dioxide (TiO2) nano-rods[46]. Karreman et al used an integrated laser scanning fluorescence microscopy and transmission EM instrument, in combination with a fluorogenic probe reaction, to map the locationof Brønsted acid domains within the heterogeneous structuralcontext of fluid catalytic cracking particles. Their integrated
approach allowed changes in chemical and functional properties of the heterogeneous catalysts to be tracked over time,
revealing how regions with changing levels of catalytic activity could be assigned to structural defects generated by aging
induced degradation of the zeolite material [47].

Figure 7. (A) When studying catalyst materials, optical microscopy
can be used to visualize chemical events, e.g. using fluorogenic
reagents (top left). By carefully tuning reaction conditions, the locationsof chemical events can even be mapped on a particle with nanometer
accuracy (bottom and right left). Unfortunately, the diffraction limit
allows only a very limited amount to be derived from the particle itself(right), making it hard to correlate chemical reactivity with nanoscale
structural features. Reproduced from [42] with permission of The
Royal Society of Chemistry. (B) The combination of SRF imaging
and EM allows individual chemical events to be correlated withultrastructure at the single particle level. Reprinted with permission
from [48]. Copyright 2017 American Chemical Society.

intercrystal intergrowths or post-synthesis modifications thatmodulate intra-crystalline diffusion or active site accessibility as
major determinants of overall performance [48, 49]. However,catalyst particles are 3D entities. Whereas confocal microscopy
and certain wide-field SRF modalities offer inherent 3D imaging
capabilities, scanning EM (SEM) imaging typically yields a 2D
view of the outer surface of the sample, preventing visualizationof internal defects and structural features [50]. While focusedion beam milling can reveal the internal structure of individual
catalyst particles [50], this approach does not scale well towards
EM imaging of larger particle volumes.
Currently, studies on catalytic systems typically follow a consecutive approach where in situ fluorescence imaging of a sample
is followed by ex situ EM imaging. While integrated CLEM systems exist, and are commercially available, further adaptations
might be needed to allow truly in situ observation of relevant
systems under ambient conditions. Reliable image registration
can be achieved using either fiducials or the intrinsic photoluminescence or cathode-luminescence to determine the coordinatetransformation between the SEM and fluorescence data34.
CLEM can also be applied to novel and often chemically diverse groups of materials such as, among many others,

Current and future challenges. CLEM studies on commerciallyapplied zeolite catalysts revealed pronounced particle-to-particle
heterogeneities, and underscore the importance of defect-rich34

13

https://github.com/KrisJanssen/QuickCLEM.

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

organometal halide perovskites. Featuring striking electricaland optical properties, perovskites are prime candidates for thedevelopment of next generation solar cells and optoelectronic
devices [43, 51, 52]. The morphology of individual domains
within condensed perovskite phases is one of the key factors
determining the generation, transport, and trapping of charges[53], particularly at domain boundaries. Moreover, phase stability and domain formation in these materials is subject to various
environmental factors such as oxygen and moisture, temperature,pressure and light irradiation, rendering these materials highly
dynamic. To understand how perovskite composition and grainmorphology influence the fate of the photogenerated charge
carriers and to ultimately predict the performance of these mat­erials in downstream applications, physicochemical and in situ
structural analysis with high spatio-temporal resolution at thesingle particle level is required. This requires the expansion ofexisting CLEM capabilities towards highly multimodal analysis
e.g.via energy dispersive x-ray spectroscopy elemental mappingor Raman scattering based imaging of chemical signatures.
Advances in science and technology to meet challenges.

Recent improvements in EM instrumentation and computational methods allow matching 3D imaging capabilities of fluorescence microscopy. In electron tomography (ET), a series ofprojection images are collected at different angles of incidenceof the e-beam using a high tilt sample holder (figure 8) [47].
A high-resolution 3D representation of the sample is obtained
computationally (figure 8 (a)) [54]. ET can be applied analytically, revealing information on the distribution of heavy elements as well as quantitatively, e.g. mapping the network of pore
structures in nano-sized zeolites (figure 8 (b)) [54].
Specialized liquid cells can be used to enable simultaneouslight irradiation and EM imaging of samples under (near) ambient, in situ conditions. Using this approach, the photocatalytic
activity of ZnO crystals at the crystal facet-level could be deduced[55]. Besides determining the link between photocatalytic performance and crystal facet expression, the influence of defects andstructural imperfections was found to be non-negligible [56].
CLEM can be extended beyond the visualization of emissive species or chemical events. Multimodal approaches combining spectral information with luminescence lifetime analysis
and localization of luminescence events reveal the dynamics ofcharge trapping sites, in response to the exposure of organometal halide perovskites to different atmospheric conditions,created in an environmental SEM [52]. Image correlation
approaches for sub-diffraction limited fluorescence imaging
can be leveraged to examine the temporal variations of luminescence intensity in CLEM which might further aid the understanding of charge carrier behaviour in complex materials [43].
In general, the potential effect of high energy electron
beams on sample systems should not be neglected. Indeed,work performed in our group indicates that the photocatalytic
performance can be affected [55], or that structural degradation of ZSM-22 crystals might occur under harsh EM imaging conditions. However, modern EM equipment with biased
stages can help to minimize these effects. The ability of theelectron beam to locally generate electric fields in materials

Figure 8. (A) The principle of ET. (B) ET allows the open (green)and closed (orange) mesopore volume in a zeolite Y crystal to bequantified. Reprinted from [54], Copyright 2015, with permission
from Elsevier.

can also be exploited to trigger certain phenomena. Recent
results on perovskites have revealed how such directional
electric fields lead to different degradation pathways compared to the light induced degradation [51].
Concluding remarks. From this short perspective, it should
become clear that with the accelerating pace of new developments in the field of correlative instrumentation, CLEM itself
might soon become a concept that is far too limited to capture therichness and depth of information that can be obtained in addition to functional and structural imaging. Multimodal correlative
analysis will prove essential to meet the demands of future applications in every sector of human endeavour, from catalysis andthe production of green chemicals to energy production and storage, as well as in bio(medical) research. In each of these fields, aproper understanding of nanoscale phenomena is essential.

Acknowledgments
We acknowledge financial support from the Research
Foundation-Flanders (FWO, Grant Nos. G.0197.11,
G.0962.13, G.0B39.15, ZW15_09 GOH6316N, postdoctoral fellowship to HY, ED and KPFJ), KU Leuven Research
Fund (C14/15/053 and IDO/12/020), the Flemish government
through long term structural funding Methusalem (CASAS2,
Meth/15/04), the Hercules foundation (HER/11/14) andthe EC through the Marie Curie ITN project iSwitch(GA-642196) and the ERC project LIGHT (GA-307523).
14

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Fluorescence and electron microscopy ofmembrane proteins within intact cells in liquid

the electron microscope, and allowing imaging with STEM
through the liquid water layer [58]. TEM is also an option,
but requires the samples to be thinner than 1 µm for nanometer resolution. Liquid flow can be initiated, helping to reduceradiation damage. The inherent cell thickness and bulging
of the chamber in the vacuum can make it challenging tokeep the liquid layer  ⩽5 µm, as needed for high resolution.
Typically, fixed cells are imaged, but with this method, cells
alive at the onset of EM can also be studied [63]. Secondly,
environmental scanning EM (ESEM) is an option for imaging cells covered under a thin liquid layer maintained in a
wet environment (figure 9 (b)). Using STEM detection [64]a resolution of ~3 nm is achieved [61] (figure 10). ESEM
allows the fastest analysis protocol, which is particularly
useful for studies involving many tens of cells. A drawback,on account of the lower electron energy of typically 30 keV
versus 200 keV of regular STEM, is a lower spatial resolution and the inability to image through thicker cell regions. A
third method involves the coverage of cells with an ultra-thinfoil, composed of graphene sheets [65], closely fitting the cell
contours, thus behaving like a flexible wrapping film [66]. It
allows the imaging of the cells in liquid with 200–300 keV
at the highest possible resolution. A current challenge is thecleanliness of the graphene, as it is often contaminated withsmall, electron-dense residues from the production. A fourthalternative, not using STEM detection, is the combination ofback scatter electron detection in SEM with LM [67, 68]. The
latter method does not provide as high a resolution as STEM
but allows the largest flexibility in the biological experiment,
because the cells are imaged in larger liquid enclosures [68]and even directly in cell culture dishes [67].
Of key importance for all liquid-phase EM methods, istheir further optimization, and in particular the developmentof additional labels to cover a wide range of membrane protein
related questions. The concept of liquid-phase EM represents
a paradigm shift; it has to be clear that the aim is not to provide
nanometer-scale information about the cellular ultrastructureor the protein structure as conventional EM does, although
this would be possible for imaging thin cellular regions. The
main value of this novel technology lies in its focus on biological and biomedical questions related to targeted membraneprotein complexes, not addressable with other methods.

Diana B Peckys1 and Niels de Jonge2,3
1

Faculty of Medicine, Saarland University, 66421 Homburg,
Germany2
INM—Leibniz Institute for New Materials, 66123
Saarbrücken, Germany
3
Department of Physics, Saarland University, 66123
Saarbrücken, Germany
Status. Electron microscopy (EM) of liquid specimens isincreasingly popular in materials science, chemistry, biology,
and other fields [57] to solve a wide range of so far unanswerable questions. In the life sciences, liquid-phase EM is mainly
used as analytical method for studying membrane proteins inmammalian cells that are kept intact and in their native liquid
environment [58, 59]. The highest resolution is obtained withscanning transmission EM (STEM). The principle relies on
the atomic number (Z) contrast of STEM, and allows detectionof specifically bound small probes, consisting of small binding peptides, or peptide tags, and nanoparticles, within several
micrometers of liquid thickness and with a spatial resolution of 1–3 nm. Imaging the locations of individual subunitsof macromolecular complexes is thus possible allowing, forexample, to determine the stoichiometry of a protein complex.
Another option is to study high-Z biological materials in cells
such as magnetite magnetosomes [60]. The unique feature isthe combination of the EM-range high spatial resolution, withthe capability to study whole cells in liquid, while avoiding
laborious preparation or destruction through sectioning or
rupture. Liquid-phase EM can easily be combined with lightmicroscopy (LM) to analyse protein expression levels andsubcellular localization via fluorescence microscopy at thesingle-cell level (figure 9), thereby addressing heterogeneityin cell populations [61]. In addition, LM prior to EM makes
it easy to navigate to cellular regions of interest during EM.
Liquid-phase EM adds a unique level of analyticalcharacterization, as it gives quantitative information at a­single-molecule and single-cell level, about the locations
and functional state(s) of the studied proteins (figure 10).
Commonly used biochemical techniques rely on extracted
material from many cells, and can thus not provide information about localization. The required resolution for direct
imaging of single subunits of protein complexes in intact cells
is not achieved by super-resolution fluorescence techniques[62]. Other indirect optical techniques, such as Förster resonance energy transfer (FRET) and fluorescence cross correlation spectr­oscopy (FCCS), have their own specific limitations,such as the imposed restriction in the FRET distance (4–8 nm),which is insufficient for large protein complexes, and the needfor very low expression levels in FCCS [61].

Advances in science and technology to meet challenges. As

with many pioneering techniques, it takes time for the researchcommunity to adapt liquid-phase EM into the common analytical research practices. More publications are needed demonstrating its capabilities for studying biological questions. Partof the problem is that biological research groups, with experience in and access to electron microscopes, are typicallyinterested in high-resolution information about the structureof cells or proteins, not optimal for liquid-phase EM. On theother hand, groups with questions on membrane proteins thatwould be suitable for this new technique, usually are experts
in methods for molecular biology and biochemistry, and may
not have frequent access to EM facilities that allow experimenting with this new technique. To date, only a few groups
use liquid-phase EM to study actual biological questions, forCurrent and future challenges. Liquid-phase EM of cells

comes in four different ‘flavours’, all combinable with correlative LM. Firstly, cells in liquid can be enclosed in a
microfluidic chamber, sealing them against the vacuum of15

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 10. Liquid-phase ESEM-STEM image of a membraneregion of a breast cancer cell showing the locations of HER2
receptors labelled with QDs. The overlay shows molecular modelsof the HER receptor with label consisting of an Affibody molecule(blue) coupled to a streptavidin-coated QD. The labels are attached
to a monomer (left) or a homodimer (right). From [61]. Reprinted
with permission from AAAS.

limit for reproductive cell death. Even bacteria do not reproduce and are thus not alive according to the definition oflife, even after exposure to such low doses. Yet, it is possible
to study cells alive at the onset of imaging taking snapshot
images of selected locations [63, 68] and it would perhaps be
doable to record time lapse image series capturing biochemical processes.

Figure 9. Correlative microscopy of membrane proteins in wholecells in liquid. (A) Cells are grown on a silicon nitride (SiN)
membrane, supported by a silicon chip. A microchip with quantum
dot (QD) labelled and fixed cells is positioned upside down in liquid
in a glass-bottom dish. Fluorescence imaging is performed with anoil immersion lens on an inverted microscope. (B) For ESEM, thesame microchip is positioned upright on a cooled stage and kept
in a saturated water vapour atmosphere. The cells are covered witha thin layer of water. Contrast is obtained on the QDs attached tomembrane proteins using the STEM detector. From [61]. Reprinted
with permission from AAAS.

Concluding remarks. By offering unprecedented spatial res-olution for the study of labelled subunits of macromolecularcomplexes in eukaryotic cells in their native liquid environ­ment, liquid-phase EM opens an entirely new field for biological EM. The acquisition of data from many tens and even
hundreds of cells in a series of experiments, comprising manytens of thousands of single-molecule data, can quantitatively
address urgent problems such as cell heterogeneity in cancer. Series of whole-cell experiments can be carried out forvarying experimental conditions, while still obtaining singlemolecule information of endogenous proteins. Correlative
fluorescence microscopy and liquid-phase EM are on their
way to develop into a key microscopy methodology for future
biological research.

example, [61, 67]. To increase its application in life sciences,
several technical issues are currently being optimized. For
instance, the nanoparticle labels used so far are larger thanthe proteins under examination, and it is expected that a size
reduction would lead to an enhanced labelling efficiency, anda higher precision in determining subunit locations. To gain
information about the cellular ultrastructure in thin cellularregions, it should be feasible to use liquid STEM tomography,
but this is yet to be demonstrated. Here, a limitation might
be imposed by the available tilting angle, therefore low-angle
tomography methods would have to be adapted. Although it
would be desirable to use liquid-phase EM for live cell observations of processes at high resolution, this option is highly
challenging and it requires careful control experiments. The
reason is the extraordinarily large radiation intensity inherent to EM, even at the lowest possible doses of 0.1 e−1 Å−2,
which is already three orders of magnitude above the dose

Acknowledgments
Ulrike Korf, David Piston, and Stefan Wiemann for discussion,
and E Arzt for his support through INM. Research supportedby the Deutsche Forschungsgemeinschaft SFB1027 (C7).

16

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Correlative light and volume electron microscopy

thereby opens the road to literally infer kinetic information to
EM images, allowing the for­mulation of a whole new array ofquestions that can be addressed with this powerful technology.

Nalan Liv and Judith Klumperman

Section Cell Biology, Center for Molecular Medicine,
University Medical Center Utrecht, Utrecht University,
Heidelberglaan 100, 3584CX Utrecht, Netherlands

Current and future challenges. Even though the tools

required for volume-CLEM are now available, some challenges still prevent wide-spread routine application. Sample
preparation for CLEM and volume-EM are both far from trivial and have evolved mostly independently from each other.
Sample preparation approaches for volume-CLEM should
therefore balance the requirements of both, and often require
extensive testing of protocol variables for each sample type.
In addition, a major challenge of volume-EM approaches is toperform immuno-labelling, the most widely used method forprotein localization studies. Most volume-EM methods, however, are not compatible with immunolabelling throughout the
sample. Pre-embedding labellings require permeabilization ofcells, compromising cellular morphology. Array tomography,
allowing the immunolabelling of an array of serial sections,
could become the unique volume-EM method for protein
localization studies in 3D, but is currently hampered by a low
throughput in the sequential analysis and correlation of constitutive sections. Because of the limited immunolabelling possibility in volume-CLEM, the identification of a discrete pointin 3D also becomes a significant challenge. The localizationand correlation of molecules in 3D between LM and volumeEM relies mainly on fluorescent tags, and this makes theaccurate correlation of 3D LM and EM data sets very crucial
for volume-CLEM. A high correlation precision is currently
achieved for 2D CLEM applications (sub 10 nm precision correlation), but still constitutes a challenge in the z-axis, becauseof the approximately 100-fold resolution mismatch between
LM and EM datasets in the third dimension.
Structure function studies of proteins or protein complexes
require quantitative imaging approaches, which is dependent on the number and volume of the sample sets that are
imaged. Therefore, higher throughput and more automatized
LM and EM instrumentation compatible with volume-CLEM
is urgently needed. Recent studies on connectomics provide
very promising examples of extended data collection [83].
Automation of instruments generates immense amounts ofimaging data, and raises the problem of storage and subsequent
analysis. It is becoming routine to collect raw images sizing up
to hundreds of gigabytes per day, and therefore, novel strategies should be considered to manage this data flood. Moreover,processing of the collected image sets, like alignment of stacks,
correlation of LM to EM and segmentation 3D rendering ofstructures, possesses another major challenge, and highlights
the need for automation not only in instrumentation, but also inimage processing and analyses. Automatized image processing
and segmentation is one of the most exciting aspects of volume-CLEM, and will enable fast and user independent quantification (e.g. volume, membrane interactions) of automaticallysegmented structures of interest in significant sample sizes.

Status. Correlative light and electron microscopy (CLEM)

infers molecular information to an ultrastructure by uniting
data from light microscopy (LM) and electron microscopy(EM). In the past decade, CLEM techniques have been usedmostly to identify rare or transient cellular events by LM, andrelate this to the underlying structural and cellular context
by EM. Hence, CLEM has become instrumental in furthering our understanding of molecular functions in cells. So far,most CLEM applications use 2D transmission EM sections. A
drawback of these approaches is that they have limited application in the z-axis, hampering the possibility to obtain 3D
images [69]. Correlative light and volume electron microscopy (volume-CLEM) broadens the applications of CLEM
to understand the function of molecules in the structural context of the 3D organization of a cell, at different organization
scales: single organelles, tissues, whole cells, and eventuallyorgans and organisms.
Recently developed volume-EM techniques that provide
ultrastructural information in 3D include: serial blockface SEM[70], where the blockface is imaged repetitively after sectioningby an in situ ultramicrotome; focused ion beam SEM (FIB-SEM)[71], where an ion beam removes slices from the blockface; andarray tomography [72], in which the serial sections from theblock are placed as an array on a substrate for large area imaging
in the x,y and z direction. In all cases, consecutive sections, or the
fresh revealed surfaces of the sample block, build up a 3D stackof images, from which the volume can be reconstructed.
Volume-CLEM approaches add LM to these EM techniques, in order to facilitate the identification of a region ofinterest (ROI), or to provide 3D structural context informationof a fluorescently localized molecule. While adding uniqueand essential information at the z-level, volume-CLEM also
poses a significant challenge over 2D CLEM: i.e. to identifiy
the same region imaged in LM back in EM in 3D. The limited
correlation accuracy in z has restricted the wide applicabilityof volume-CLEM studies so far. However, time invested byimaging scientists to develop novel workflows is paying off,
and uniting CLEM with volume-EM has now been successfully applied to visualize zebrafish development [73], brain
tissue [74–77], and cellular processes like bacterial and viral
infection mechanisms [78, 79].
CLEM is most rewarding when used to relate dynamic
events, only visible in live cells, to an ultrastructure (livecell CLEM) [80, 81]. The advancements in volume-EM haveenabled the evolution of live cell-CLEM to live cell-volumeCLEM, providing a direct link between live cell dynamics,
functional imaging (proteins at work) and 3D ultrastructure [16,82]. The unique combination of parameters provided by live
­cell-volume-CLEM can be integrated on the single cell or evensingle organelle level (figure 11). Live-cell-volume-CLEM

Advances in science and technology to meet challenges.

Recent innovative developments in automated volume-EMare significantly reducing the user interaction times needed to17Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 11. Volume-CLEM providing a direct link between live cell dynamics and 3D ultrastructure on the single organelle level [82]. (A)Schematic representation of the complete live-cell fluorescence to volume-EM workflow. (B) As an example, a fluorescence image of a
LAMP-1-GFP transfected cell, incubated with Dextran-Alexa568 as an endocytic marker. (C) The cell was imaged live for several minutes,
followed by in situ fixation. Stills show the LAMP-1-GFP spots (spot 1, 2) during 142 s of imaging. After fixation the cell is stained,
embedded in resin, and imaged in FIB-SEM. (D) Shows the slices on all three viewing axes (XZ/XY/YZ) of the reconstructed FIB-SEM
dataset containing the live-cell ROI ((B), (C) red square). Both spots 1 and 2 are classified as late endosomes based on their high number of
intraluminal vesicles. (E) FIB-SEM segmentation and 3D reconstructions of spots 1 and 2; the organelles imaged in live-cell fluorescencemicroscopy (1,2) were segmented and correlated with reference LM data.

obtain large volumes of high-quality data. These include themulti beam SEM system of Zeiss which uses 61 e-beams forparallel image acquisition [84], the multi energy deconvolution imaging strategy from FEI/ Thermo Fisher, which createsisotropic imaging data by ascending the e-beam energies totarget different depths [85], and the impressive 196 multibeam system with a probe size of 1 nm [86]. These instrumentation developments contribute to the generation of immense18Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

magnitudes of structural information within considerably lesstime, and pave the way to relating structural information to
direct measures of function, as said, a major application forvolume-CLEM.
Functional information can be best linked to structure
when molecules are first imaged at work (live-cell imaging), and then localized in the context of their 3D ultrastructure. Considering the limited immunolabelling possibilities
of volume-CLEM, the precision with which the molecules
can be localized on the ultrastructure is the main challengeof volume-CLEM sample preparation. Genetically encoded
peroxidase derivatives chemically convert diaminobenzidine
and produce an electron dense reaction product. When fusedwith a fluorescent protein these constitute a promising molecular localization technique compatible with both LM and volume-EM datasets. Interestingly, recent studies showed thatmolecular localization studies do not necessarily depend onelectron dense stains, but can be accomplished with bimodal
fiducials [87]. Volume-CLEM approaches would greatly benefit from sample preparations protocols with 3D fiducials(preferably intracellular), as well as algorithms to accurately
correlate LM and EM datasets in 3D. Therefore, the development of endocytic tracers which can be used as fiducials
and 3D correlation software, is highly warranted. Similarprobes could be used for correlation in cryo-EM approaches.
Another approach for molecular localization is preserving
the fluorescent signal after resin embedding. In particular,
the development of fixation-resistant fluorescent probes (e.g.
meos4b), in combination with LR-white or HM20 acrylic
resins, is providing encouraging results. Although the sensitivity of this approach is currently modest, because of areduction in signal due to embedding, the application isespecially interesting for recently developed integrated room
temper­ature CLEM microscopes [88, 89]. The developmentsof a next generation of integrated volume-CLEM microscopes (e.g. LM in a FIB-SEM), or their combination with
serial section based approaches, is very favorable for fast andaccurate correlative imaging for 3D analysis of cells, tissues
and model organisms.
Storage and analysis of the precious volume-CLEM data
is not a trivial process, and poses a challenge on the community to create common solutions. Creating public microscopyarchives for volume-CLEM, e.g. electron microscopy public
image archive [90], would not only provide a solution for raw
data storage, but also enable scientists to analyse the deposited data and extract the information relevant for their specificscientific questions. Developments in automated image processing, e.g. semi-or fully-automatized stack alignment andautomated segmentation of volume-EM datasets, are urgently

needed and in the focus of attention. Full automation of thewhole pipeline, i.e. collection of LM and volume-EM data,
correlation, stack alignment, reconstruction and final segmentation, is the final goal and being addressed by the boomingfield dedicated to EM image analysis.
Cryo-volume-CLEM approaches, in which LM and volume-EM are performed at cryogenic temperatures, avoid
chemical fixation, staining, and dehydration, and allow visualization of molecular structures with Angstrom resolution. The
booming cryo-EM field would greatly benefit from cryo-volume-CLEM to identify ROIs and infer molecular information
to EM images. The challenges to address, in order to exploit
the full potential of cryo—CLEM, are both on the cryo-LM
and cryo-volume-EM sides. These include instrumental considerations about the z-resolution of cryo-LM, adaptation ofsuper-resolution techniques, as well as understanding the spectral characteristics of fluorophores which are different at cryo
conditions than at RT [91]. On the volume-EM side, the recent
advances in cryo-EM of macromolecules are revolutionizing
structural biology. However, there is yet no solution for EM
imaging of extended volumes in cryo conditions, and the true
potential of cryo-volume-EM application at the cellular levelremains to be explored. Cryo-volume-CLEM approaches will
greatly benefit, when both the LM and EM techniques matureand their resolutions in all 3D will match better.
Concluding remarks. Biological systems encompass manyorganization scales, i.e. molecules, organelles, cells, tissues
and organs, in healthy and disease conditions. The complexityof these systems is based on delicate balances, which depend
on much more than a simple sum of the characteristics of small
entities. It is increasingly appreciated that biological systems
should be studied and evaluated in higher content, and byinvestigating the system as a whole, rather than a reductionist
approach focused on simplification. Investigating the massive
dynamic interplay between millions of molecules even in asingle cell, requires high-content and high-resolution imaging
techniques, which can relate molecular dynamics to the ultrastructure. This makes volume-CLEM approaches indispensable for paving the way to novel biological breakthroughs, inwhich the identification of tagged macromolecules, analysis of
their functions within living cells and tissues, and their ultrastructure, are assessed in an automatized quantitative manner.

Acknowledgments
We thank the members of our group, especially Job Fermie, for
input and discussions.

19Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Force spectroscopy and single molecule fluorescence microscopy: unpacking single viruses

resolution, the sample has to be robust enough to withstandthe forces exerted by the scanning tip and remain stationary
while acquiring the data. Both aspects seriously limit the
number of biological processes that can be studied with AFM.
Additionally, AFM imaging has a rather poor discriminating
power—most proteins will look like nanometer sized blobs,
which makes their identification difficult in more complex
samples. Although impressive progress has been made withhigh-speed AFM towards the observation of moving objects,
it remains challenging to visualize biological processes when
their dynamics are not slowed down. Imaging of freely diffusing specimens is impossible and will only lead to an increased
noise level as such objects intermittently collide with the tip or
cantilever. To overcome these limitations, it is a logical step tocombine AFM with optical microscopy. To this end, AFMs arealmost routinely placed on top of inverted optical microscopes
to perform correlative studies, i.e. where AFM and optical data
are obtained one after the other. Performing AFM and fluorescence simultaneously requires a drastic isolation of the AFM
from mechanical vibrations that are introduced by the added
microscope parts. Especially during force spectroscopy experiments, nanometer vibrations will translate in force fluctuations of tens to hundreds of piconewton. Typically, the AFM
heads on inverted optical microscopes are fairly large, whichleads to a large mechanical loop to the sample. This increases
drift and lowers the resonance frequencies of the system. The
resulting vulnerability to vibrations hampers nanometer andpiconewton resolution. In addition, vibrations and thermal
drift are induced by illumination sources and cameras which
often contain a cooling fan. In part, these noise issues can becontrolled by placing the instrument on an active isolationtable within an acoustic and temperature controlled enclosure.
This, however, leads to complicated and costly instrumentsthat remain out of reach for most, except some specializedlaboratories. Even so, their performance remains inferior to
the stand-alone AFMs they are based on.
At the latest, when AFM and optical microscopy are combined, it becomes clear that both techniques have contradicting demands on sample preparation. AFM is performed on
samples immobilized at the surface, at concentrations of tens
per square micrometer, while florescence microscopy does not
require immobilization and is generally performed at lower
sample concentrations.

Iwan A T Schaap1 and P J de Pablo2
1

SmarAct GmbH, Schütte-Lanz-Str. 9, D-26135 Oldenburg,
Germany2
Dpto. Física de la Materia Condensada Universidad
Autónoma de Madrid 28049-Madrid and Instituo de Física dela Materia Condensada IFIMAC, Universidad Autónoma de
Madrid 28049, Madrid, Spain
Status. The emergence of atomic force microscopy (AFM)

has made it possible to observe biological structures with sizes
that are well beyond the limits of conventional optical microscopy. Because the technique works in liquids, samples canbe studied under conditions that are representative for their
physiological environment. Furthermore, AFM based force
spectroscopy is intensively used for the mechanical manipulation of biological matter. Initial experiments, originatingfrom physics laboratories, demonstrated that proteins shells
like microtubules and viruses exhibit a clear elastic responseat small deformations followed by breakage events at higher
forces [92, 93]. The relevance of such mechanical characterization is evidenced by the acceptance by the biological andbiomedical communities and has, for example, led to the discovery of clear correlations between the mechanical state of avirus and its infectivity [94, 95]. Further interest for mechanical measurements is generated by the evolving insight thatforces play critical roles in biological processes, ranging fromthe unpacking of viruses at nuclear pore complexes to controlling cell differentiation [96, 97].
An essential condition for AFM is the immobilization of thesample to the surface, which prevents the observation of diffusive processes occurring in solution. This is unfortunate because
the majority of biological processes rely on diffusion, and thus
remain out of reach for AFM. To take the application of AFM to
the next level, it is essential to surpass this limitation. AFM force
spectroscopy has been combined with fluorescence microscopy
to study protein unfolding [98] and we have recently combined
single molecule fluorescence with AFM imaging and force
spectroscopy, to track the release of viral DNA upon mechanical stimulation of a 100 nm small virus, thus combining thecomplementary benefits of each of the techniques (see figure12) [99]. Only when the simultaneous operation of AFM based
force spectroscopy and single molecule fluorescence microscopy finds its way out of the physics laboratories, will it be able
to fulfil its full potential for life science research.

Advances in science and technology to meet challenges. Allof the technology needed to address the aforementioned challenges is available. Below, we list the main requirements toachieve simultaneous AFM based force spectroscopy and single molecule fluorescence, for studying bio-macromoleculesin the context of life science, with sizes below the diffraction
limit.

Current and future challenges. When operated in liquid under

low force conditions, AFM can resolve biomolecules withnanometer resolution. In addition, by pulling or pushing with
the AFM tip, the mechanical response of the sample can beprobed, which provides a method to better understand its structural organization. This combination of n­ anometer-resolution
imaging and piconewton-resolution force spectroscopy makes
AFM a seemingly ideal tool for studying biological processes at a single protein level. However, to obtain such high(1)	Integrated instruments with less noise. (i) To reduce theeffects of the mechanical loop between tip and sample,
it is key to use a stable design (thermally and mechanically), which can be achieved by reducing the size of the
AFM as much as possible [100]. (ii) In combined instruments, multiple piezo scanners and positioning stages
20

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 12. (A) A schematic of a simultaneous force spectroscopy and single molecule fluorescence experiment on a virus. An AFM

tip is used for both imaging and manipulation purposes at the nanoscale. The molecular response is simultaneously tracked by imaging
fluorescent markers (green) that are excited by an evanescent wave (blue). The microscope objective is shown in grey. (B) An example froma virus before and after mechanical manipulation. Top row: AFM scans (200  ×  200 nm), showing the disassembly of a virus capsid. Bottom
row: fluorescence images (4  ×  4 µm) showing the release of YOYO-1 labelled DNA. The DNA remains invisible in the AFM scans because
it is not immobilized to the surface.

are required to move the AFM, with respect to both thesample and the optical microscope. These need to bearranged and controlled such that the positioners thatare not used for AFM will also not introduce vibrations
during AFM measurements. A notorious example is theclosed-loop piezo objective scanner used for focussingthat can couple vibrations into the sample through theimmersion oil. iii) Optical imaging does not requirephysical contact between the sample and the microscope
and this should be exploited. In a minimal configuration,
only the oil-immersion objective and a short-range focussing rack need to be implemented in the AFM stage. Allother components (lenses, filters, cameras, lasers) can bemechanically uncoupled from the AFM sample stage [99].
(2)	Limit artefacts caused by the AFM tip in the opticaldetection volume. During force spectroscopy measurements, the AFM tip and cantilever will be very close to
the sample and thus scatter light, which can dramatically increase the fluorescence background signal [101].
Fluorescence excitation based on a total internal reflection scheme is ideal, as this limits the exited region toabout 100 nm from the microscope cover slip and thus
leaves the AFM cantilever and most of the tip in the dark.
Confocal microscopy is, for this reason, much harder tocombine with AFM.
(3)	Fluorescence super-resolution techniques. The resolution
gap between AFM and optical microscopy is more than
two orders of magnitude. Confocal microscopy based
super-resolution techniques are, for the reason mentioned
under the previous point, less suitable, but techniques
based on the accurate localization of single fluorophores[102] such as stochastic optical reconstruction microscopy(STORM) [103] can help to further increase the information that can be obtained during the measurements.
A simple approach to obtain fluorescence information

beyond the diffraction limit during force spectroscopy
was applied during the unpacking of single viruses [99].
More advanced approaches like STORM will offer higher
spatial resolution but at the cost of temporal resolution.
(4)	Sample preparation. Studying biological processes withdiffusive factors requires strategies in which only a subsetof the components are immobilized to the surface, whileother components remain free to diffuse [104]. A specificfluorescence response can be achieved with fluorophores
that respond to mechanical or optical events. This can
include Förster resonance energy transfer pairs and caged
compounds, but also intercalating dyes that gain access to
DNA upon mechanical stimulation [99].
Concluding remarks. AFM can greatly benefit from the integration with the current state-of-the-art optical microscopy
approaches. Optical detection is especially suited for identification purposes and for those parts of a sample that are soft or
highly mobile. AFM imaging provides the nanometer resolution context for the localized mechanical manipulation of thesample. Once the simultaneous combination of AFM and single molecule fluorescence has reached the maturity requiredfor its employment by biological laboratories, its application
is not limited to studying relatively simple systems, like thecargo release by viruses. Indeed, an opportunity will be created to study multifold biological processes that are incompatible with current AFM approaches.

Acknowledgments
P J de Pablo acknowledges projects FIS2014-59562-R,
FIS2015-71108-REDT, Fundación BBVA and the ‘Maríade Maeztu’ Program for Units of Excellence in R&D(MDM-2014-0377).

21

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

High-speed atomic force microscopy and lightmicroscopy

and encapsulated substrate proteins, (iv) detecting minutechanges in protein molecules that are only detectable withfluorescence energy transfer, and (v) observing protein molecules under an external force applied via optical tweezers.
The purposes (i) and (ii), not requiring simultaneous HSAFM/LM observations, are already accomplished [107, 111].
A hybrid HS-AFM/fluorescence microscopy (including totalinternal reflection fluorescence microscopy) is already used(figures 13(c) and (d)) together with a wide-area scanner thatfacilitates a post-experiment positional correlation of AFM/fluorescence microscopy images [111]. Far-field super-resolution
fluorescence microscopy, such as stimulated emission depletion microscopy and single-molecule localization microscopy,
is already combined with conventional AFM [112]. Combining
with HS-AFM will be readily accomplished.
The purposes (iii)–(v) impose challenges. In current
HS-AFM, the sample stage is scanned. When this system iscombined with LM, the LM image oscillates, and more seriously, the optical path of LM is blocked by the cantilever.
In tip-scan HS-AFM, the laser beam focused on the cantilever has to be steered quickly to track the cantilever motion,
at least in the lateral direction. A 2D mirror tilter has been
recently developed, which can perform this tracking at 1 kHz,up to 20 µm [111].
One of the other challenges is to correlate between HS-AFM/
LM images captured in synchrony. This is because of a large
difference in the field of view; the scan range in HS-AFM imaging of protein molecules is typically ~100  ×  100 nm2, while thefield of view in far-field LM is ~30  ×  30 µm2. Moreover, thespatial resolution gap between HS-AFM (2–3 nm) and conventional LM (200–300 nm) also makes this correlation invest­
igation difficult. Besides these issues, the susceptibility of thecantilever to strong pulsed laser illumination in LM, or to highpower YAG-laser illumination in optical tweezers, is anotherissue to be considered in the hybrid system.

Toshio Ando

Nano Life Science Institute (WPI-NanoLSI), Kanazawa
University, Kanazawa, Japan
Status. Structural biology has solved the atomic structure ofmany proteins, but all revealed are static snapshots. Single-molecule biophysics has recorded the dynamic action of proteins but
only in an indirect way of observing optical markers attached tothe molecules. Therefore, it has been a desire to acquire a means
to directly observe protein molecules in dynamic action. Highspeed atomic force microscopy (HS-AFM) was developed tomake this long-awaited observation possible. The state-of-theart HS-AFM system allows us to image protein molecules at
10–20 frames s−1 (fps), without disturbing their function. Its
spatial resolution is typically 2–3 nm and ~0.15 nm in the lateral and vertical directions, respectively. Despite this moderatespatiotemporal resolution, HS-AFM holds a unique and pivotal
position among various tools for biological research. It is only
HS-AFM that allows the simultaneous assessment of structureand dynamics of single molecules.
A variety of purified protein systems have been successfully
visualized by HS-AFM [105]. Even protein molecules on thesurfaces of live bacteria [105] and nuclei [106] have also beenvisualized. Moreover, the development of fast, wide-area scanners has given HS-AFM the capability to visualize live eukaryotic cells [107]. The HS-AFM movies of proteins have provided
new mechanistic insights that could not have been reached inother ways, as exemplified by the discoveries of an adenosine
triphosphate (ATP)-independent generation of intramoleculartension and powerstroke in myosin V [108], rotary propagationof chemical and conformational states in rotor-less F1-ATPase(figure 13(a)) [109], unidirectional motion of cellulases hydrolysing cellulose fibers [105], and the formation of ESCRT III
spiral filaments with unnatural curvatures, leading to membrane
buckling, budding and abscission (figure 13(b)) [110]. Besides,
HS-AFM was demonstrated to be able to visualize the very thinand highly flexible structure of intrinsically dis­ordered proteins[105]. As illustrated by these examples, the improvement fromstatic/indirect to dynamic/direct observation can advance in
depth and detail our understanding of how proteins function.
And yet, HS-AFM still has plenty of room for improvement and functional expansion towards various directions,one of which is hybrid HS-AFM/light microscopy (LM). LM,
particularly fluorescence microscopy, provides information
complementary to that acquirable with HS-AFM.

Advances in science and technology to meet challenges. HS-

AFM can visualize dynamic conformational changes of protein molecules induced by ligand binding/release. However,
it is often not easy to judge whether or not visualized conformational changes are really those induced by ligand binding/release. Simultaneous HS-AFM/fluorescence microscopy
observations facilitate this judgement, when fluorescentligands are available. However, correlation identification
between HS-AFM/fluorescence images is also required.
The achievable single-nanometer resolution of far-field
super-resolution fluorescence microscopy is incompatible
with high temporal resolution. Moreover, most of the techniques rely on strong pulsed laser illumination. These features
are inadequate for materializing simultaneous HS-AFM/fluorescence microscopy imaging.
Near-field scanning optical microscopy (NSOM) also beats
the diffraction limit and therefore can be a candidate to be used in
the HS-AFM/fluorescence microscopy hybrid system. NSOM
uses either aperture probes for local illumination or apertureless probes for local optical detection/illumination [113]. AFM/
NSOM hybrid systems already exist, where the AFM cantilever(metal-coated) tip is also used as an aperture (figure 14(a)) or

Current and future challenges. This hybrid microscopy isrequired for the following purposes: (i) positioning the AFM
tip on a region of interest within a large object (e.g. eukaryotic
cells) visualized with LM, (ii) localizing specific molecules
labelled with fluorophores within an overlaid topography
image of complicated biological architectures, (iii) observing interaction of HS-AFM-visualized protein molecules withother molecules invisible for HS-AFM, such as small ligands
22Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 13. (a) HS-AFM images of α3β3 subcomplex of F1-ATPase undergoing conformational changes in the presence of adenosine
triphosphate (ATP). The height of a nucleotide-free β subunit is larger than those containing ATP or adenosine diphosphate. The highest
pixel positions marked with red dots shift counterclockwise. At 1.44 s, the α subunit adjacent to the nucleotide-free β subunit appears higher
than the β subunit. Imaging rate, 12.5 fps. (b) HS-AFM images showing spiral filament formation by polymerization of the ESCRT-III
protein Snf7 on a supported lipid membrane. Imaging rate, 3 fps. (c), (d) Simultaneously captured HS-AFM (c) and total internal reflection
fluorescence microscopy (d) images of Cy3-labeled chitinase A (arrows) moving unidirectionally on a chitin microfibril. Imaging rate, 3 fps.

Figure 14. Various configurations of cantilever-based NSOM. (a) Aperture NSOM relying on a specially designed cantilever chip. The

tip has a small optical opening at the apex, where a localized evanescent light is produced. (b) Aperture-less NSOM based on a metalcoated cantilever tip that acts as a local electromagnetic antenna (plasmon resonator). (c) NSOM combining a small aperture with anelectromagnetic nano-antenna. The nano-antenna is a metal particle with the shape of a bow-tie, rod, sphere or dumb-bell.

aperture-less (figure 14(b)) probe for NSOM. The simultaneous, pixel-by-pixel acquisition of the optical and AFM signals,
at nearly identical sample positions, facilitates finding a correlation between the two overlaid images. In aperture NSOM,
the spatial resolution achieved so far is 50–100 nm, while it is10–25 nm in aperture-less NSOM. However, in aperture-less
NSOM, the optical contrast is deteriorated by background signals produced by the far-field or evanescent illumination of awide area. Recently, this problem has been overcome by combining an aperture probe with a metal (mainly gold or silver)
particle, having special shapes, as an optical nano-antenna(figure 14(c)); the strong plasmon resonances localized aroundthe particle are used as a local illuminator [114].
To combine this plasmonic aperture NSOM with HS-AFM,
micro/nano-fabrication techniques have to be established, toproduce short cantilevers with both an aperture probe andnano-antenna. Moreover, a fabrication technique to produce nano-antennas under precise size/shape control is also
required. When an HS-AFM/NSOM hybrid system is used toobserve the interaction between a protein molecule and a fluorescently labelled ligand (e.g. ATP) molecule, high spatial resolution, 5–10 nm, will often be required for NSOM, especially23

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

for homo-oligomeric proteins. This high spatial resolution isalready reported but has not become routine. This is primarily because of a lack of techniques for the mass production of
NSOM probes with identical properties.
HS-AFM visualization of single protein molecules under
external force can provide new mechanistic insights into theirdynamic processes, such as unfolding or refolding processes,
and processes undergone by force-responsive/force-generating proteins (e.g. mechano-responsive receptors/channels,
mechano-sensor kinases, cell adhesive proteins, and motor
proteins including DNA helicases and viral DNA packaging
proteins). Rather than constructing an HS-AFM/optical tweezers system, its operation is challenging because the operator has to handle HS-AFM, LM and optical tweezers, while
avoiding cantilever-YAG laser beam contact. Therefore, anautomation control system is important in facilitating thiscomplicated operation.

Concluding remarks. Truly useful and innovative correlation

microscopy of HS-AFM/LM should allow simultaneous fastrecording of the two modalities of images with well-defined
pixel-by-pixel correlation. To materialize this imaging, higher
resolution NSOM has to be routinely achieved. At the sametime, micro/nano-fabrication technology has to be establishedto produce small cantilevers with a nano-illuminator for plasmonic NSOM. When the objects to be visualized by NSOMin this correlation microscopy are small ligand molecules, wewill encounter an additional challenge to synthesize fluorescent functional ligands. The HS-AFM/optical tweezers hybrid
system will provide a new opportunity to study the unfolding/refolding processes of proteins and directly observe protein
molecules responding to an external force, exerted at a specificlocus in a given direction. Although it is difficult to foresee thefuture progress in these hybrid systems, their materialization
will surely lead to new important discoveries of proteins.

24Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Traction force microscopy

fibers can generate force through clutch-like mechanisms in afibrous matrix, using 3D TFM [122]. Only through TFM has it
become evident how important physical forces are for biological systems, e.g. for cell adhesion, migration, differentiationand fate (‘mechanobiology’).

Ulrich S Schwarz1 and Christian Franck2
1

Institute for Theoretical Physics and BioQuant, Heidelberg
University, Heidelberg, Germany
2
Department of Mechanical Engineering, University of
Wisconsin-Madison, 1513 University Ave, Madison, WI 53706,
United States of America

Current and future challenges. TFM has certainly excelled atproviding high-resolution detail on cellular force generationin single cells in 2D, and more recently in 2D monolayers and3D, but in order to keep pace with the ever-increasingly complex biological questions in mechanobiology, it now has toevolve along many different directions at the same time. As the
biological applications become more demanding and sophisticated, opportunities exist to further refine the resolution andintegration capabilities of TFM into quantitative and correlative biophysical frameworks. At the molecular level, we needincreased integration with imaging and high throughput data.
On the imaging side, this may comprise close integration withsuper-resolution and light sheet-based imaging techniques ofcell adhesions, the cytoskeleton, the nucleus and the extracellular matrix, as well as fluorescent sensors reporting local
activation of regulatory molecules (such as the small GTPases
Rho and Rac) and forces on single molecules. Regarding highthroughput advances, TFM has already been combined with
RNA-interference in a high throughput manner, and it could
also be combined with other high throughput techniques, suchas mass spectrometry, CRISPR/CAS9 gene editing, or single
cell sequencing. At the tissue level, we expect a move towards
more medically relevant multicellular systems, e.g. clusters ofcancer cells moving through fibrous tissue or organoids withspecific functions grown from induced pluripotent stem cells.
This will create interdisciplinary opportunities for biologists,
physicists and engineers to work side-by-side to address these
exciting new challenges. At the same time, however, this also
creates the need for rigorous quantitative benchmarking andvalidation data sets, to evaluate and standardize the evergrowing arsenal of TFM and TFM-like methodologies thatwill emerge in the process. As it is common in bioinformatics
and image processing, there is also a need to write and maintain publicly available TFM-software that can be used easily
by a large user community. Finally, the handling and processing of large amounts of multispectral, spatial and time-lapse
data will need to be addressed, especially for high-throughput
or clinical investigations. The use of machine-learning andartificial intelligence algorithms may provide unique opportunities to link up with TFM, to address both multi-modal imaging and high-throughput data challenges.

Status. Traction force microscopy (TFM) was originallydeveloped in the late 1990s as a technical procedure to convert
the deformations of a flat elastic cell culture substrate (typically made from polyacrylamide gel) into an estimate of cellular traction forces, using the equations of elasticity theory[115]. Because cells on flat substrates spread very thin andpull on their substrate, mainly tangentially, it is often sufficient to restrict the force reconstruction to two dimensions(‘2D TFM’) (figure 15). Early efforts in the field focused on
improving the reliability and efficiency of this method, forexample, by performing the calculations in Fourier space,introducing regularization techniques from control theory, andby improving the image processing methods required to track
substrate deformations [126].
Around the year 2010, TFM started to explore new areas.
The first studies that showed reconstructed traction forces not
only for single cells, but also for cell ensembles of different
sizes, from cell doublets to endothelial monolayers (‘mono­layer
stress microscopy’) [117]. The most important aspect of thesestudies was that cells could now balance their forces not only
against the substrate, but also against neighboring cells. Aroundthe same time, TFM leaped into the third dimension, both byreconstructing vertical cell forces on flat substrates (‘2.5 TFM’),and by considering cells fully embedded in three-dimensionalhydrogels (‘3D TFM’) (figure 16) [118]. To date, some of themost physiologically advanced 3D TFM techniques featurecollagen gels, which have important medical applications, suchas wound healing or cancer metastasis. For cells in such 3D
fibrous environments, methods were developed to either establish an appropriate material model [119] or to formulate a TFMprocedure that does not require a material law [120].
Based on these advances, at present, various TFM form­
ulations exist for reconstructing cellular force fields fromnanometer to millimeter length scales, using either closedform analytical solutions for relatively simple material platforms, or finite element and hybrid formulations accounting
for more complex material interactions. The main goal andintent of TFM is the correlation of physical forces with cellular, often structural, data, ranging from the size of individualadhesion sites through the spatial organization of the cytoskeleton, to the migration characteristics of a whole cell. Theseobservations have spurred the development of hybridized
advanced mathematical models that are capable of providing more detailed structural information than TFM experiments alone (‘model-based TFM’). Such approaches havebeen used to estimate the forces inside individual stress fibers
via 2D TFM [121], and to understand how individual stress

Advances in science and technology to meet challenges. Currently, a large effort has been invested in

improving the procedures of classical TFM, using concepts
and methods from physics, engineering and materials science[115–117]. First, one requires an accurate methodology formeasuring cell-generated substrate displacement fields on aroutine basis, but with high precision [120]. While there exist
many popular choices of single particle tracking, particle
image velocimetry and optical flow schemes, they might fail25

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 15. 2D TFM. (A) Movement of marker beads in the flat elastic substrate are converted into an estimate of the cellular forces atadhesion sites. (B) An adherent U2OS-cell has many adhesion sites (marked by paxillin) and actin-based stress fibers. (C) Image processing
is used to track substrate displacement. (D) Elasticity theory is used to calculate the traction stresses. (E) The image data can be used with amechanical model for the cell to estimate forces in individual stress fibers. Reproduced from [121]. CC BY 4.0.

in the case of large deformations. A recurrent issue is also theneed to track these deformations in relation to a stress-freereference configuration. Recently, a method has been developed to circumvent this need by printing a marker system
onto the undeformed substrate (‘reference free TFM’) [123].
Next, we need to improve on the accuracy of our physical material (constitutive) description of the substrate fromwhich the displacement and traction fields are deduced, bothin 2D and 3D. Even to date, most TFM investigations haverelied upon the use of polyacrylamide gels, which possess
excellent idealized homogeneous and elastic material behaviour. Yet, as tissue culture models move more towards physiologically-realistic extracellular matrix design, or even into
real tissue, TFM invest­igations will need to extend their own
framework to account for more complex material descriptions[119]. This, in turn, will require the development of advanced
experimental techniques for providing accurate descriptionsof the local, as opposed to bulk, mechanical properties of theextracellular matrix surrounding the cells. Finally, an appropriate physical model for converting cellular displacements
into surface tractions and forces is needed [120]. The development of tightly integrated experimental-theoretical frameworks [121, 122] will allow us to resolve more geometricallycomplex boundary value problems of cell–matrix and cell–
cell interactions, e.g. clusters of cancer cells migrating andgenerating force in 3D.
Going beyond the classical formulation of TFM, the field
is wide open for innovative approaches to reconstruct cellular forces from the deformation of a calibrated deformation
sensor. For example, two interesting techniques have recently
emerged to measure cell forces from podosomes, which
are vertically organized adhesion sites used for invasion byimmune, bone and cancer cells. In the first technique (‘protrusion force microscopy’), an atomic force microscope isused to measure podosome-generated bumps on a thin elastic
membrane [124]. In the second technique (‘interference stressmicroscopy’), an optical interferometer is used to detect cellforces to the top surface of an elastic resonator cavity [125].
Both techniques do not require any measurement of a stressfree reference state and track deformations directly with very
high resolution. These examples show that creative innovations can be used to fill gaps in the classical methodology, toanswer very specific biological questions.
Concluding remarks. While the original TFM methodologies have allowed us to appreciate the significance of physical26

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 16. 3D TFM. (A) Movement of marker beads in a 3D soft matrix are converted into an estimate of the 3D cellular displacements

forces at adhesion sites. (B) A fully embedded human neutrophil (red) inside a collagen-I matrix (blue). (C) Image processing is used to
track 3D matrix displacements. (D) 3D surface projections onto the cell surface for deduction of matrix displacements and strains. Cellularsurface tractions can be deduced from (D) if matrix properties are known. Reproduced with permission from [120].

forces in cell biology, more advanced formulations will beneeded to keep pace with the rapid development in the life sciences, materials design, tissue engineering and imaging technologies. As biological investigations open up new frontiersin revealing complex biological behaviour, quantitative TFM
approaches will be required to correlate the wealth of 2D and
3D fluorescent data to physical interactions inside, outside
and among cells. This, in turn, provides a tremendous opportunity for experimental and modelling communities to cometogether and develop new methods, frameworks and models toquantify and correlate the biological world.

Acknowledgments
U S S is a member of the cluster of excellence CellNetworks
and of the Interdisciplinary Center for Scientific Computing(IWR) at Heidelberg.

27

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Super-resolved traction force microscopy

is subject to aberrations induced by the mismatch between
the refractive index of the gel and that of the immersion
medium [116, 126]. Improvements in aberration correction
using, for instance, specialized objectives with refractiveindex mismatch corrections, as well as using adaptive optics,
would reduce the impact of these effects and would result in
an improved spatial resolution, possibly along all three spatial dimensions, allowing even higher bead densities to be
used and maximizing the accuracy of the TFM and STED
instrumentation.
While the localization accuracy of STFM is theor­eticallynot limited, for any given experiment, the sampling of the displacement field must meet the Nyquist criteria i.e. the spatialsampling frequency of the displacement field must be twicethat of any feature that may be resolved in the displacement
field [127]. This restricts the density of fiducial markers thatmust be high enough to reflect the complexity of the traction field that is applied by the cell. If the bead density is too
low, areas of the gel will move without being reported by any
bead movement, and the traction information will be lost.
Conversely, if the bead density is too high, information will
also be lost, as a result of the point spread functions of eachindividual bead overlapping with those nearby, making theirrelative displacements hard to recover accurately. Further, themaximal bead density within the gel must also be balancedwith the ability to track the beads and the fluorescence lightsensitivity of the biological specimen. Consequently, the quality and robustness of STFM experiments could be improvedby better preparation of the gels with precise, regular, and predetermined bead concentrations e.g. using substrate printing
systems.
STFM precision is also affected by the acquisition durationof the fiducial marker positions. Only sufficiently fast acquisition times allow accurate tracking of the bead positions overtime, which directly depends on the scanning speed of the
microscope laser beam. Faster scanning beams, therefore, canfurther improve the quality of STFM experiments.
The resolution performance in super-resolution microscopy is profoundly related to the properties of the fluorescent dyes. Superior fiducial markers, using fluorescent dyeswith improved photo-stability, switching performance andquant­um efficiencies, reduce the light illumination of thecells and thus prolong the total time of STFM experiments.
In this way, the accuracy and the total time period of STFM
experiments, and thus the fluorescence light sensitivity of thebiological specimen, can be improved by brighter and morestable fiducial markers.
The spatial resolution of STFM also depends on thereconstruction method used to recover the displacement of
the beads from the fluorescent images [126, 127]. The mostcommon methods of extracting bead displacements are those
based on single particle tracking, where each individual beadmust be localized, and those methods based on statisticalcompariso­ns of fluorescent images, such as particle image
velocimetry (PIV). To this end, the image is divided into agrid, and each grid element is spatially correlated between
frames to assess the degree of movement. PIV does not
require localization of each bead but is limited spatially by

H Colin-York1, C Eggeling1,2,3 and M Fritzsche1,2
1

MRC Human Immunology Unit, Weatherall Institute of
Molecular Medicine, University of Oxford, Headley Way,
OX3 9DS Oxford, United Kingdom
2Kennedy Institute for Rheumatology, University of Oxford,
Oxford OX3 7LF Oxford, United Kingdom
3
Institute of Applied Optics, Friedrich Schiller University,
Jena, Germany
4
Leibniz Institute of Photonic Technology (IPHT), Jena,
Germany
Status. Super-resolved traction force microscopy (TFM)

has the potential to transform live-cell mechanical force
measurements [126, 127]. Conventional TFM is perhaps themost commonly used technique for mechanical probing incells, owing to the simplicity of its implementation on mostlaser-scanning confocal microscopes. In TFM experiments,
cells adhere to the protein functionalized surface of a thin
(20–30 µm) elastic gel [116]. Within the gel, immobilised
fluorescent beads serve as fiducial markers, and imaging of
the bead positions over time during the application of cellular tractions, allows the elastic displacement of the gel to bequantified. Combining the displacement measurements withthe mechanical properties of the gel allows the forces appliedby the cell to be recovered. The greatest shortcoming of TFM
is the limited sensitivity due to the finite density of the fiducial
markers, at which a displacement field can be sampled [126].
Consequently, the resolution limit of conventional fluorescent
microscopes imposes a fundamental limit on the length scaleof mechanical probing [128].
To overcome this challenge, we recently improved the spatial resolution and accuracy of TFM using ­super-resolution
optical stimulated-emission-depletion (STED) instead ofconfocal or wide-field microscopy [126, 127]. The increased
spatial resolution of super resolution TFM (STFM) allows agreater than five-fold higher sampling of the forces generated
by the cell than conventional TFM, leading to more accuratequantification of cellular tractions (figure 17).
STFM naturally suffers from the restrictions inherent to
all TFM and STED experiments, but it is likely to becomea standard-tool for the quantifications of mechanical force
production in living cells, because the resolution of STFM istheor­etically not limited, as it scales with the applied STED
power [129]. Ultimately, improving the spatio-temporal sampling of force measurements allows the quantification of
mechanical force production in unprecedented detail, providing biomedical research with a state-of-the-art force probing
technology to unravel the molecular mechanisms underlying
mechanical force production in living cells.
Current and future challenges. Several important challenges

must be addressed to further improve the (S)TFM protocol.
Advances can be achieved in the TFM and STED instrumentation, as well as in the fluorescent dyes of the fiducial markers.
The nature of the (S)TFM setup requires all imaging to
be done at the top surface of the gel, meaning that imaging
28

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 17. STFM applied to HeLa cell focal adhesions. (a), (b) Traction magnitude calculated from the measured confocal (a) and STED(b) recordings of the bead displacements. Scale bars, 2 µm.

the size of the grid elements required to give accurate correlations. Hence, the development of more advanced reconstitution algorithms could improve the quality and robustness of
STFM experiments.

Concluding remarks. STFM imaging is established as avaluable tool for biological research and its application must
be extended to a wider range of scientific questions. The
development of STFM will involve the implementation of
improved fluorescent dyes with increased photo-stability,
allowing a reduction in photo-toxicity. In addition, newmicroscope technologies will be necessary, as extensions tomore complex biological specimens will benefit from further
improvements in the spatial and temporal resolution, as wellas the ability to image thicker specimens. Combining STFM
with high-performance objectives for deep-substrate imaging
and/or the application of adaptive optics, holds a significant
potential benefit for mechanical probing in immunology, biophysics, and other areas. Further, the increasingly improved
usability and availability of super-resolution systems will
reduce the need for external technical support, thus facilitating the potential of STFM to become the standard tool formechanical force probing in the field of biomedical research.

Advances in science and technology to meet challenges.

Emerging technical advances in optical super-resolution
microscopy and the design of fluorescence dyes present theopportunity of establishing high-performance mechanicalprobing as a day-to-day toolbox, in the near future.
To this end, the recent development of 3D-STED[120] allows an even greater sampling for STFM of theforces perpend­icular to the gel surface, in the same waythat 2D-STED can increase the accuracy of the tangential
force reconstruction. TFM has previously been successfully
extended to measure 3D bead displacement, generated bycells within 3D elastic environments [120].
Fluorescent beads are bright and photo-stable, but are limited by photo-bleaching at high frame-rates or long acquisition
times. Conventional beads could be replaced by customdesigned fluorescent nano-materials, such as nano-diamondswith luminescent properties, e.g. nitrogen-vacancy centersthat possess good bio-compatibility, high photo-stability, andfacile surface functionality [130].
Acquisition of the fiducial markers is limited by therecording duration of the observation region, which could be
improved by parallel scanning of multiple regions of interests,
or by wide-field based illumination of the complete field ofview through, for instance, structured illumination [131].

Acknowledgments
We greatly acknowledge support from the Wolfson Imaging
Centre and funding by the Medical Research Council(grant No. MC_UU_12010/unit programme G0902418 and
MC_UU_12025), MRC/BBSRC/EPSRC (grant No. MR/
K01577X/1), Wellcome Trust (grant reference 104924/14/Z/14
and Strategic Award 091911 (Micron)), Wolfson Foundation,
and Oxford internal funding (EPA Cephalosporin Fund, John
Fell Fund).

29

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Electrophysiology on lipid bilayers combined withfluorescence imaging and spectroscopy

in vitro system, artificial bilayers can reproduce the full range
physico-chemical properties of the biological membranes, andare thus an adequate environment for the functional reconstitution of membrane proteins. The expansion of the classical bilayer
technique, where ion channels are analysed by electrical single
channel measurements, in combination with the nanoscopic fluorescence techniques (microscopy/spectroscopy) in HLBs, decisively extends the possibilities of this in vitro-system. (i) Free
standing HLBS and partially supported HLBs in combination
with high-resolution fluorescence techniques allow to examine;

Richard Wagner1, Satya Prathyusha Bhamidimarri1, Niklas
Brending2 and Mathias Winterhalter1
1

Department of Life Sciences & Chemistry, Jacobs University,
Bremen, Germany
2
Ionovation GmbH, Osnabrück, Germany
Status. Classical planar lipid bilayers containing reconstituted

ion channels, transporters and pumps have already served forlong time as a well-defined model system for high-resolution
electrophysiological investigations of ion channel and large
pore forming solute channels (porins) functions. More recently,
suitably designed microchips containing free-standing horizontally oriented bilayers (HLBs), allow the combination of electrophysiology with high-resolution nanoscopic fluorescence
techniques down to the single-molecule level. Combining
simultaneous single-molecule fluorescence measurements ofmembrane protein conformational changes with high-resolution
electrophysiology, has been generally regarded as a promising
tool to gain insights into the structure–function relationships ofion channels at the single molecule level [135]. As proposedalmost 20 years ago, the holy grail of membrane channel studies
is to produce an atomic scale movie of an ion channel or large
porin-like channel solute at work, simultaneously observing
conformational and electrical or solute property changes, as ionsor uncharged solutes flow through the protein [136]. Regardlessof the early identification of this challenge, one has to admit thatup to now only a few publications employing directly correlated
simultaneous fluorescence and electrical recordings on single
ion channels appeared in high-ranking journals [134]. HLBs,
in combination with nanoscopic fluorescence techniques,
were also successfully employed to probe physico-chemical
properties of bilayers and the topology of membrane proteins[134, 137]. Obtaining a molecular movie of a single transporter slow turnover action would require HLBs containing
these transporters, while solely using nanoscopic fluorescencemicroscopy [28] to monitor transport and simultaneous changes
in the protein. One attempt to follow membrane transport ofuncharged solutes at a single molecule level in HLBs, is the
optical single transporter recording approach, using confocal
microscopy and HLB-arrays [138]. The combination of highresolution fluorescence and electrical recording allows themonitoring of the transition of water soluble toxins, which formhigh conducting membrane pores upon transition from solution
into the membrane [132]. Corre­spondingly, it should be possible to monitor the transition of water soluble receptor-cargocomplexes into the membrane, thereby forming membranepores [139]. Finally, fluorescent ligand binding combined withsingle-channel recording, is aimed at providing mechanistic
insight into the behavior of ligand-gated channels [28].

(i)	the properties of lipid mixtures forming the bilayer membrane.
(ii)	the topology and oligomeric states of membrane proteins(iii)	the transition of water-soluble proteins forming membranepores upon transition into the membranes (pore-forming
toxins, receptor/cargo complexes).
(iv)	Also, ligand gating of ion channels can be investigatedusing fluorescent effectors, and electrical single channel
recording can give molecular details on ligand binding
affinities and how binding is transmitted to channel gating.
(v)	Low rate transport of uncharged metabolite solutes bylarge pore forming channels like the classical bacterial
porins can be investigated using electrical and confocal
fluorescence recording with HLBs.
One recent example for this is given in figure 19, whichshows the HLB containing multiple copies of LamB, a substrate specific E. coli channel protein, which contains specificmaltose binding sites and is part of the bacteria-specific maltodextrin transport pathway. We investigated the specific bindingof a fluorescent MDP composed of maltohexaose, carrying aperylene fluorophore (MDP-1). Through fluorescence correlation spectroscopy (FCS) measurements along different pointsof the z-axis, we could characterize three different mobility
classes of MDP-1, with τ(dif ) ∼= 0.2 ms (P1) in solution toτ(dif ) ∼= 25 ms (P2) close to the bilayer and τ(dif ) ∼= 62 ms at
the bilayer bound to LamB (see figure 19(C)). The results
show also that MDP-1 is not permeable through LamB.
Concluding remarks. HLBs, in combination with the single-

molecule techniques of the electrical single channel recording
and nanoscopic fluorescence microscopy/spectroscopy, offer
great potential with regard to their ability to resolve biologicalprocesses in vitro down to the molecular level of single steps.
Despite this, few publications on the use of these combined
techniques are found in the literature. In our opinion, this ismainly due to the high technical requirements of the individual disciplines. So far only very few robust HLB-systems withpossible access to the solution on both sides are commerciallyavailable. In order to facilitate the combined technique, we
believe it is necessary to develop fully integrated HLB systems
in which arrays of bilayers are automatically generated. Thesearray chips with automated liquid handling must be individually, electrically addressable. Such an automatic HLB system,transferred into a suitable environment with state of the art
measuring technology for nanoscopic fluorescence techniques(microscopy/spectroscopy) and electrical single/multichannel
techniques, could develop the full potential of the technology.

Current and future challenges. Single molecule methods provide a unique insight into the function of biological systems,
compared to macroscopic average values. They are primarily
able to measure the distribution of the biomolecule behaviour orproperties and not just their ensemble mean values. As a robust
30Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 18. Principal setup of a horizontal lipid bilayer (HLB) combined with a confocal scanning spectrometer allowing high resolution

fluorescence imaging and spectroscopy [132–134].

Figure 19. (A) shows a stack of confocal x-z scans through a LamB containing bilayer (POPC/POPE/POPS 8:1:1) after addition of 60 nM
MDP-1 (cis side, buffer symmetrical (cis/trans) 1 M KCl 10 mM HEPES pH 7.2) at t  =  0 recorded with time intervals of Δt  =  2 minfor each following x-z scan (total distance per scan 100 µm). (B) Recorded fluorescence intensity along the HLB z-coordinate. (B)Corresponding molecular brightness along the z-direction. (C)–(E) FCS measurements along the HLB z-coordinate. (F) Electrical single
channel recording from a bilayer revealing that MDP-1 induces channel blocking only from the cis site. As shown by sequential confocal
z-scans of the HLB MDP-1 when added to the cis HLB-compartment binds in a time dependent course to the bilayer (A) and (B) andremained stably bound to the membrane after repeated careful perfusion of the cis compartment (not shown). Intensity profile along thez-direction of the HLB (D) indicates that MDP-1 is stable bound to LamB in the bilayer which was confirmed by competition with nonlabelled MDP which released the bound MDP-1 from the membrane (not shown).

Medicines Initiatives Joint Undertaking under Grant Agreement
No. 115525, resources that are composed of financial contrib­
ution from the European Union’s seventh framework program(FP7/2007–2013) and EFPIA companies in kind contribution.

Acknowledgments
The research was conducted as part of the TRANSLOCATION
consortium and has received support from the Innovative31

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Voltage-clamp and patch-clamp fluorometry:studying ion channels and transporters with lightcontributes to the spectroscopic response, is identical to the
population contributing to the electrophysiological response.
However, in practice there might be unlabeled proteins contributing to the current and labeled proteins being electricallysilent. The second precondition for a useful correlation is toperfectly synchronize spectroscopic and electrophysiologicaldata.
As in other live-cell imaging approaches, several sourcesof background fluorescence can disturb spectroscopic recordings. One major source is autofluorescence, the natural emission of light by biological components. Except for approaches
in which genetically encoded fluorophores are used, background fluorescence can additionally arise from (1) unspecificbinding of fluorophores to molecules nearby the proteins ofinterest, (2) unspecific binding to equipment surfaces like
patch pipettes, and (3) unbound fluorophores in the surrounding bath solution. Importantly, negative control recordings todetermine background intensity have to be performed undervoltage-clamped conditions, because fluorophore binding toproteins and surfaces might be voltage-dependent.
However, one of the biggest challenges in VCF/PCF, as inlive-cell imaging in general, is to find a fluorophore whose
physical and chemical features match the experimental needs,
and which does not affect the protein behavior when being
attached to it. To this aim, it is required to (1) identify themost suitable probe (excitation and emission wavelength, sizeand chemical structure, pH dependence, environment depend­ence, unspecific binding), (2) check for the optimal positionin the protein, (3) check for the best tagging strategy (posttranslational labeling, genetically encoded protein dyes, fluorescent unnatural amino acids), and (4) take into account sizeand chemical structure of linkers between fluorophores andproteins.

Jana Kusch1 and Giovanni Zifarelli2
1

University Hospital Jena, Jena, Germany
Department of Physiology, Anatomy and Genetics, Universityof Oxford, Oxford, United Kingdom

2

Status. To gain a deep understanding of the complex

dynamic behavior of proteins, structural and functional dataneed to be integrated. For ion channels and ion transporters,functional information can be collected with an extraordinaryhigh time-resolution by voltage-clamp and patch-clamp techniques. These approaches, with their numerous variations,revolutionized ion channel/transporter research significantly.
Nevertheless, conclusions about structural rearrangements
within a protein, dynamic protein interactions or changes
in protein mobility, can only be drawn indirectly. Structural
methods, like x-ray crystallography, nuclear magnetic resonance or cryo-electron microscopy, on the other hand, produce a wealth of structural information, with a resolution
down to the atomic level. However, these approaches provide
only static snapshots of highly dynamic processes and in mostcases, the functional state associated with a given structure is
not even fully defined.
To overcome these limitations, classical electrophysiological approaches were combined with different kinds of spectroscopic techniques. Thus, ion channel/transporter function canbe monitored simultaneously with conformational changes,
changes in protein or subunit interactions, ligand binding
events, or protein mobility, giving the possibility to directly
correlate these functional and structural/spatial parameters.
The first step in this direction was made in the 1990s, throughthe establishment of the so-called voltage-clamp fluorometry(VCF), that employed either the two-electrode voltage clamp[140] or the cut-open technique [141] to investigate ion channels in whole Xenopus laevis oocytes. Typically, in VCF,
labeling of the protein is carried out by site-directed sulfhydryl-modification of cysteine residues, but more and more frequently by fluorescent unnatural amino acids. Inspired by theprinciples of VCF, several types of patch-clamp fluorometry(PCF) approaches were designed, combining different electrophysiological and spectroscopic tools, currently only forion channels [142]. The various components reported so far
are summarized in figure 20 [143].
Despite the results already achieved, both VCF and PCF
are still under continuous development. Whereas up to now,
mostly in vitro experiments on protein ensembles were performed, further advances might set additional foci on singlechannel and in vivo approaches.

Advances in science and technology to meet challenges. As

mentioned before, VCF and PCF are still under continuous
development, particularly with respect to their spectroscopic
components. Advances in both labeling and microscopy technologies are required.
Regarding labeling technologies, we expect that the rapidly
progressing developments in the field of channel/transporter
research will demand a parallel development of new and rather
custom-tailored fluorophores. Like for other live cell imaging
approaches, the challenge is to develop fluorescent molecules
which are not only small, bright and photo-stable, but which
also have a low cellular toxicity, offer a high signal-to-noise
ratio, and do not interfere with the proteins of interest. For
post-translational labeling, fast labeling kinetics and, in thecase of intracellular targets, good membrane permeability isdesired. Probe selection and design can be simplified because
there is only a small number of ‘core’ fluorophores giving rise
to a large ensemble of fluorescent probes by attaching various
reacting groups, substrate moieties, chelating components,and other chemical entities to these core molecules [144].
This continuous development of new fluorophores and theincreasing variety of biological systems used in PCF require aparalleled development of gentle chemical strategies, to mask
unspecific binding sites in the sample. Preventing unspecific

Current and future challenges. The core of VCF/PCF

approaches, is to correlate two sets of different types of data(ion current and fluorescence changes), which are collectedusing different types of recording devices in the same experiment. In this context, a major challenge in most of the PCF
approaches is to ensure that the protein population, which
32Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 20. Overview of the main electrophysiological andspectroscopic tools combined in PCF. The high variabilityof PCF, given by the number of possible combinations ofelectrophysiological and spectroscopic tools, makes it a potent
approach for studying ion channels, thus overcoming the limitationsof the individual techniques. Reprinted from [143], Copyright 2014,
with permission from Elsevier.

binding to equipment surfaces like glass substrates, demands
new passivating materials which do not interfere with opticaltransmission or cell attachment.
To tackle the problem of autofluorescence, the strategy ofchoice is avoidance, demanding advances in labeling and microscopy technologies, which go hand in hand. This should provide
better fluorophores with long excitation wavelengths, as well as
optimized excitation and emission filters, and allow for improved
dual wavelength correction or linear unmixing settings.
In VCF/PCF, as in fluorescence microscopy in general, there
is an inherent trade-off between spatial resolution, time resolution and phototoxicity. Despite huge progress in the development of high- and super-resolution microscopy over the lastyears, further efforts are needed to get rid of sample fixation,paving the way for monitoring highly dynamic processes ofproteins or protein domains without motion-induced blur. On
the part of microscopical components, further optimization of
objective lenses, filter sets, and detectors will help to acquire
high resolution images without cell or protein damaging.

Figure 21. Three examples illustrating the wide variety of combining
electrophysiological and spectroscopic approaches. (A) VCF with
Xenopus laevis oocytes expressing voltage-dependent potassiumchannels (KV1.2/2.1 chimera). Fluorescence signals arising fromthe incorporated unnatural amino acid Anap (red traces) and gating
currents (black traces) were recorded in response to depolarizing
voltage jumps. Reproduced with permission from [145]. (B) PCF
combining single-particle tracking and whole-cell patch clamp.
Neurons expressing P2X2FLAG–YFP receptors were labeledwith QDs and imaged over time with epifluorescence optics(black trajectories). At the same time, whole-cell voltage-clamp
electrophysiology (green trace) was used to measure transmembranecurrents before, during and after ATP applications. Reproduced withpermission from [146]. (C) PCF combining confocal laser scanningmicroscopy and inside-out patch-clamp configuration. Patches were
excised from Xenopus laevis oocytes expressing mHCN2 channels.
Fluorescently labeled cAMP (fcAMP) was applied to study ligand
binding (green trace) and channel activation (black trace) in responseto an activating hyperpolarizing voltage jump (black triangle).
Reprinted from [147], Copyright 2010, with permission from Elsevier.

Concluding remarks. Over the last two decades, VCF and

PCF have been shown to be tremendously powerful approaches
to shed light on the dynamic behavior of ion channel proteins.
Essentially, there seems to be no limit to the imaginable combinations of spectroscopic and electrophysiological tools (figure 21). However, with the exception of a few studies dealing
with single-channel recordings, up to now, the focus was clearlyon the investigation of protein ensembles in vitro, monitoring alarge number of channels, often several hundreds or thousands
at the same time. Further efforts are expected to establish thetechnical conditions to facilitate more single-channel andin vivo experiments.
So far, in contrast to VCF, PCF has not been applied toion transporters, most probably because of relatively low cur­rent amplitudes, particularly in cell patches. New strategies toincrease protein expression levels and new recording devices
with higher sensitivity might expand the applicability of PCF
to transporter proteins in future.
Acknowledgments
We thank Vasilica Nache for proofreading and commenting.
Funded by the Deutsche Forschungsgemeinschaft (DFG,
German Research Foundation)—TRR 166 project B7 to J K.
33

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Fluorescence and magnetic resonance imaging

and BOLD fMRI utilise the EPI readout sequence. The specificity of dfMRI signals has been intensely debated, as they aresuspected to be contaminated by the T2* BOLD signal [153]. A
key step in validation is to determine the specificity of dfMRI ina system that is free from blood flow interference.
Neuronal activation has been routinely measured in vitro
using brain slices and calcium-sensitive fluorophores, todetect the influx of calcium ions [154]. Therefore, a hybrid
imaging system that enables simultaneous in vitro brain slicestudies using MRI and a calcium fluorophore can be used tovalidate dfMRI. A hybrid MRI and FI system (figure 22) was
recently built using a single-sided MRI at 0.32 Tesla (T) to
test dfMRI in organotypic cortical cultures [155]. The low
magnetic field and the use of an RF surface coil provided thebenefit of an open space above the magnet to accommodate achamber with access for sample manipulation, a continuous
perfusion system to control the sample physiological state,
and the placement of optical hardware close to the sample for
efficient detection of the fluorescence signals. This system,
however, only produced MR signals (not MR images) so it
could not be used to investigate spatial resolution of the diffusion signal. The use of a low-field magnet may also limit thesensitivity of detection of the diffusion signal.

Nyoman D Kurniawan, Viha Parekh, Michael W Vogeland David C Reutens

Centre for Advanced Imaging, The University of Queensland,
Brisbane, QLD 4072, Australia
Status. Preclinical studies using in vivo magnetic resonanceimaging (MRI) often require downstream ex vivo histology
and fluorescence imaging (FI). FI microscopy is vital for thevalidation of the underlying cellular changes observed by
MRI. Currently, MRI and FI are rarely performed simultaneously, due to issues relating to optical hardware sensitivity andcompatibility within the magnetic field, differences in sample
preparations and temporal/spatial resolutions.
FI can utilise a vast array of biomarkers to study distinct
cellular processes, and discrete information can be obtained
from the same sample using reporter fluorophore molecules.
Depending on the sensitivity of the optics, FI can capture
images from micrometre to millimetre resolutions in single
cells and tissue slices in vitro [148] and in live animals [149].
Ultra-high resolution FI is mostly performed with in vitro
samples due to constraints on fluorescence labelling, opticalmagnification, depth of penetration and light scattering.
MRI detects the resonance radio frequency (RF) signals from
nuclear spins inside a strong magnetic field, and is routinely
used to obtain high-resolution images from intact living animals.
The ability of MRI to image at sub-cellular resolution, its temporal resolution and the range of available MRI biomarkers, aremore limited compared to FI. However, MRI has the advantageof yielding excellent tissue contrast without external labellingto measure biological structure and function, for example using
proton spin relaxation and microscopic water diffusion [150].

Advances in science and technology to meet challenges.

New hardware advances in MRI involving the developmentof ultra-high field magnets (17 T for preclinical MRI), parallel imaging, cryoprobes and field gradients (>1 T m−1)
are important to increase image sensitivity and resolution.
Powerful gradient coils are important to obtain strong diffusion gradients within a short echo-time and to minimise
artefacts in high field MRI. Parallel imaging and cryoprobes,
however, will be difficult to implement in a hybrid systemwith FI, due to spatial constraints within the bore of the MRI.
New advances in FI involve the development of in vivo
two-photon microscopy (TPM) [156] and ultra-sensitive
photomultiplier (PMT) charged-coupled devices, which areuseful to detect fluorescence signals from a single cell. New
calcium-fluorescence indicators with no resting signal (e.g.
Fluo4-AM) can be used to further enhance fluorescence detection upon calcium binding.
A proof-of concept hybrid platform for simultaneous imaging of a whole ex vivo mouse brain using TPM and MRI has
recently been built at 16.4 T [157]. Key aspects of this system
are the protection of the TPM module from the strong magn­
etic field, where laser excitation was directed to the sample via
a set of relay lenses and mirrors. The fluorescence emission
was relayed to the PMT module using a long light guide cable(figure 23). The TPM module produced ultra-high resolution
images at a cellular level, but only within a small field-of-view;
this was complemented by high-resolution images of the wholebrain using MRI. Unlike a hybrid system using a single-sided
low-field magnet, the space within the bore of the ultra-highfield magnet is typically very limited. Therefore, there may bechallenges in accommodating a chamber for live brain slices
and apparatus necessary to achieve a constant flow of nutrients,and obtaining images in the absence of flow artefacts.

Current and future challenges. Conventional brain functional

MRI (fMRI) indirectly detects neuronal activation by measuring the hemodynamic response given by T2* dependent blood
oxygenation-dependent (BOLD) signals. BOLD fMRI contains
some temporal delays because the hemodynamic response isdelayed by mechanisms underlying neurovascular coupling. In
addition, BOLD fMRI may depict a larger area of brain activation
as the signals can also originate from veins and capillaries that are
distal from the site of neuronal activation [151]. Therefore, thedevelopment of new fMRI methods that directly detect neuronal
activation are necessary to accurately map brain activity.
Neuronal activation involves membrane depolarisation
with the initial influx of sodium ions, calcium ions and water
being followed by the release of potassium ions into the extracellular space. The transient increase in intracellular water
results in cell swelling, in which the proportion of slow water
diffusion inside the cells increases, compared to the fast water
diffusion outside the cells. Detection of such activity-dependent swelling has been proposed as a means of direct detectionof neuronal activity using diffusion fMRI (dfMRI) [152].
Detection of pure dfMRI signals in living animals has beendifficult. To achieve adequate temporal resolution, both dfMRI

34

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 22. A platform for simultaneous in vitro fMRI and FI. (a) This system is based on an open MRI at 0.32 T (bottom table), a brainslice chamber with constant nutrient flow (middle) and a camera (top) for detection of fluorescence. (b) Two surface coils placed below andover the slices, (c) microscope image of the brain slice, (d) simulated excitation profile of the coils. [155] John Wiley & Sons. Copyright ©
2015 John Wiley & Sons, Ltd.

Figure 23. A proof-of-concept ex vivo platform for TPM and MRI at 16.4 T. The laser scanning module (upper left corner) was locatedoutside the 16.4 T magnet room and the excitation laser beam was guided into the in-MRI imaging module (centre image) using fibreoptics.
Simultaneous view of whole mouse brain using TPM (top right corner) and MRI (bottom right corner). Reproduced from [157] CC BY 4.0.

Concluding remarks. The development of the platform forsimultaneous MRI and FI acquisitions are still in its infancy.
This is an exciting multi-disciplinary field of research requiring expertise in MRI, optical FI, instrumentation and neuroscience. The development of such hybrid systems will besignificant not only for the validation of dfMRI, but also forproviding a platform to understand the basis of MRI signals
change at the cellular level.
Acknowledgments
Mr Donald Maillet for engineering expertise.
35

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Fluorescence microscopy and scanning smallangle x-ray scattering: imaging of biological cells

A subsequent study by Bernhardt et al [161] studied humanmesenchymal stem cells, and established an automated principal component analysis (PCA) of the diffraction patterns. This
allowed for the identification of the orientation of filamentous
actin, as well as the degree of anisotropy, in which the actin
inflicts on all scattering components in its surroundings. Thisapproach was extended to a full correlation of acto-myosin
fiber structures in cardiomyocytes by fluorescence microscopy and scanning SAXS [158]. Figure 25 shows the correlation of fluorescently labeled and identified stress fibers (topleft) using the filament sensor [162] and the PCA determinedorientation of the SAXS signal (top right), whereas on thebottom, the histogram of angular deviations between the two
methods is shown to center around 0° [163].
Nowadays, correlation of fluorescence and SAXS signals
can be readily achieved at a sufficient signal-to-noise-ratio
for freeze-dried and vitrified samples, as shown in [159, 160].
However, it is highly desirable to conduct such measurements in aqueous environments and ultimately probe living
cells. Towards this goal, the first SAXS recordings of living
cells have been demonstrated based on SAXS compatible cellchambers in [164], and subsequently in [165]. Importantly,
significant changes in the structure factor, due to chemical
fixation, were demonstrated in [165]. The main challenge
in live cell SAXS recordings is without a doubt the narrowwindow of allowable dose, assuring on one hand sufficientsignal—which is more difficult in the hydrated than in freezedried state—and on the other hand preventing radiation damage, which is also accentuated in the living state and aqueous
medium, due to the transport of free radicals.

Florian Rehfeldt1 and Tim Salditt2
1

University of Göttingen, Third Institute of Physics—
Biophysics, 37077, Göttingen, Germany
2
University of Göttingen, Institute for X-ray Physics, 37077
Göttingen, Germany
Status. Fluorescence microscopy has enabled a massive gainin knowledge about the localization of proteins in cells, andthereby of intracellular structures, based on selective labelingusing antibodies. Over the last two decades, super-resolution
fluorescence microscopy has helped to replace the Abbe resolution limit by the size of the fluorescent label, as the ultimate resolution limit. While nanobodies, Fab fragments, andsimilar approaches, have significantly decreased this size, allthese approaches are still relying on external labeling of thesample [28]. To overcome both the limit of label size and theconstraints associated with only detecting the labeled entities,
complementary techniques are required. To this end, small
angle x-ray scattering (SAXS) is well known to resolve soft
matter and biomolecular structure in unlabeled biomolecularsolutions and suspensions, with a resolution down to below
1 nm, as limited by the largest scattering angle. While in its
conventional form it is limited to ensemble averages, SAXS
can be extended to offer real-space resolution and hence, a
localization of the diffracting structures by combination withfocusing optics [158].
Until recently, scanning SAXS was exclusively used forstrongly scattering biomaterials such as bone or wood, andmostly with moderate x-ray focal spot sizes of several µm.
Progress in x-ray optics and detection has now made it possible to record even the rather weak SAXS signal at the level ofsingle organelles in biological cells, as was first demonstrated
in 2012, both for bacteria [159] and eukaryotic cells [160].
Scanning the sample through a micro- or nano-focused x-ray
beam, the total SAXS signal around the beam stop provides
an x-ray darkfield contrast for each pixel, which is sensitive tomolecular structures by ways of probing Fourier componentsof the electron density variations. Apart from the total scattering cross-section encoded in this darkfield SAXS signal, thediffraction pattern in each image pixel can be analyzed in viewof anisotropy, form factors and structure factors, see figure 24.

Advances in science and technology to meet challenges. Achieving SAXS for hydrated biological cells, witha signal quality that allows for quantitative analysis beyond
recording a simple darkfield, is extremely challenging. It will
require an optimization strategy to reach the required signalto-noise without (or with tolerable) radiation damage. The
x-ray photon energy has to be decreased with respect to thediscussed prior work [159–165] probably to a range between
4–6 keV, which is largely sufficient to penetrate a cell, andwhich offers an increased SAXS signal per dose [166]. This,in turn, requires improvements in sample chamber design tokeep the water layer and window materials sufficiently thin,and therefore the background low. Here, windows of crystalline freestanding thin films are best suited. Microfluidic
exchange of the (degassed) solution is required as well as highquantum efficiency in detection, and careful cleaning of thebeam from spurious SAXS background. Moreover, spatial
resolution has to be sacrificed. Sufficient SAXS quality may
be reached only for spot sizes in the micron range, but notin the nano focus range. All of these requirements are useful
but not indispensable for cryogenic (vitrified) cells. Here, theexploitable dose window is extended to 108 Gy. The challenge
here is sample preparation and transfer to a cryo-compatible
scanning stage, which is not commercially available. Contrarily, freeze-dried cells will be relatively easy to record, butof course only of limited interest.

Current and future challenges. The combination of fluorescence microscopy and SAXS, carried out on the same cell in acorrelative approach, offers a powerful new approach to gain
additional insight into the nanostructure of biological samples,by including structures which cannot be readily identified
by fluorescence markers. Image analysis of both modalities
will require automated image registry and advanced image
processing. A first correlative approach was demonstrated
by Weinhausen et al [160], recording fluorescence and x-ray
darkfield signals from the same keratin filament bundles inkeratin over-expressing human adrenal cortex carcinomaderived cells.
36Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Figure 24. Scanning x-ray microscopy using a micro- or nano-focused beam. (a) Fluorescence microscopy shows the actin fiber

distribution that is pixelwise analysed in (b). (c) Composite image of the scan area including the two rescaled eigenvectors (gray and black)from PCA. (d) Total SAXS signal provides an x-ray darkfield map and (e) anisotropy from PCA for each pixel. (f) Radial intensity profiles
can be fitted to get more information in terms of form and structure factors. Reproduced from [163]. © 2017 IOP Publishing Ltd andDeutsche Physikalische Gesellschaft. CC BY 3.0.

Ultimately, one also wants to combine super-resolution
light microscopy in situ with scanning x-ray scattering methods. However, this challenge has certain technical limits, e.g.
no current immersion optics are applicable, as the sample
needs to be kept in air for x-ray measurements. A novel setup
allowing STED microscopy, with an air objective in a horizontal geometry directly at a nano focus synchrotron beamline
suitable for scanning SAXS, is under way (Bernhardt et al,unpublished). This will also be extremely useful to shed lighton the mechanisms and constraints of radiation damage, since
the cells can be monitored by STED directly after x-ray exposure. Finally, novel analysis schemes are required to quantify
structural information recorded in a combination of real space(by scanning), as well as in reciprocal space (by recording ofdiffraction patterns). Importantly, the real space data can also
help to guide the formulation of suitable models to interpretthe scattering signal.
Concluding remarks. We briefly reviewed the current statusof correlating fluorescence microscopy and scanning x-ray
scattering on biological cells and laid out some ideas forthe future. In particular, the combination of scanning SAXS
with STED microscopy is promising to shed light on cellularcomp­onents, such as the nano structure of the cytoskeleton,
the nuclear envelope, and DNA organization. In the future,correlative approaches may help to elucidate how those structures are involved in the complex process of transcription andgene regulation. Probing the unlabeled local electron density,in addition to recording labeled biomolecules, will bring theup to now invisible structures ‘to light’.

Figure 25. Correlating fluorescence and x-ray scattering anisotropy.
(a) Most significant filaments determined by the filament sensor
from a fluorescence micrograph of neonatal rat cardiac tissue. (b)PCA-results of a nano-diffraction scan showing the order parameterΩpa (grey scale) and filament orientation θpa. (c) Histogram of
the angular deviation between both methods revealing excellent
correlation. Reproduced from [163]. © 2017 IOP Publishing Ltd
and Deutsche Physikalische Gesellschaft. CC BY 3.0.
37

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001

Acknowledgments[11] Briggman K L and Bock D D 2012 Volume electron
microscopy for neuronal circuit reconstruction Curr. Opin.
Neurobiol. 22 154–61[12] Rigort A and Plitzko J M 2015 Cryo-focused-ion-beam
applications in structural biology Arch. Biochem. Biophys.
581 122–30[13] Hauser M, Wojcik M, Kim D, Mahmoudi M, Li W and
Xu K 2017 Correlative super-resolution microscopy:
new dimensions and new opportunities Chem. Rev.
117 7428–56[14] Peddie C J, Domart M C, Snetkov X, O’Toole P, Larijani B,
Way M, Cox S and Collinson L M 2017 Correlative superresolution fluorescence and electron microscopy using
conventional fluorescent proteins in vacuo J. Struct. Biol.
199 120–31[15] Hildebrand D G C et al 2017 Whole-brain serial-section
electron microscopy in larval zebrafish Nature 545 345–9[16] Collinson L M, Carroll E C and Hoogenboom J P 2014
Correlating 3D light to 3D electron microscopy for systems
biology Curr. Opin. Biomed. Eng. 3 49–55[17] Balzarotti F, Eilers Y, Gwosch K C, Gynna A H, Westphal V,
Stefani F D, Elf J and Hell S W 2017 Nanometer resolution
imaging and tracking of fluorescent molecules withminimal photon fluxes Science 6325 606–12[18] Schnell U, Dijk F, Sjollema K A and Giepmans B N 2012
Immunolabeling artifacts and the need for live-cell imaging
Nat. Methods 2 152–8[19] de Beer M A, Kuipers J, van Bergen En Henegouwen P M P
and Giepmans B N G 2018 A small protein probe forcorrelated microscopy of endogenous proteins Histochem.
Cell Biol. 149 261–8[20] Glenn D R, Zhang H, Kasthuri N, Schalek R, Lo P K,
Trifonov A S, Park H, Lichtman J W and Walsworth R L
2012 Correlative light and electron microscopyusing cathodoluminescence from nanoparticles with
distinguishable colours Sci. Rep. 2 865[21] Somlyo A V, Shuman H and Somlyo A P 1977 Elemental
distribution in striated muscle and the effects ofhypertonicity. Electron probe analysis of cryo sections
J. Cell. Biol. 3 828–57[22] Scotuzzi M, Kuipers J, Wensveen D I, de Boer P, Hagen K C,
Hoogenboom J P and Giepmans B N 2017 Multi-color
electron microscopy by element-guided identification ofcells, organelles and molecules Sci. Rep. 7 45970[23] Leapman R D 2017 Application of EELS and EFTEM to
the life sciences enabled by the contributions of Ondrej
Krivanek Ultramicroscopy 180 180–7[24] Nisman R, Dellaire G, Ren Y, Li R and Bazett-Jones D P
2004 Application of quantum dots as probes forcorrelative fluorescence, conventional, and energyfiltered transmission electron microscopy J. Histochem.
Cytochem. 1 13–8
[25] Adams S R et al 2016 Multicolor electron microscopy for
simultaneous visualization of multiple molecular species
Cell. Chem. Biol. 11 1417–27[26] Jiang H, Kilburn M R, Decelle J and Musat N 2016
NanoSIMS chemical imaging combined with correlative
microscopy for biological sample analysis Curr. Opin.
Biotechnol. 41 130–5[27] Betzig E, Patterson G H, Sougrat R, Lindwasser O W, Olenych S,
Bonifacino J S, Davidson M W, Lippincott-Schwartz J andHess H F 2006 Imaging intracellular fluorescent proteins at
nanometer resolution Science 313 1642–5[28] Hell S W et al 2015 The 2015 super-resolution microscopy
roadmap J. Phys. D: Appl. Phys. 48 443001[29] Johnson E, Seiradake E, Jones E Y, Davis I, Grünewald K and
Kaufmann R 2015 Correlative in-resin super-resolution andelectron microscopy using standard fluorescent proteins Sci.
Rep. 5 9583

F R acknowledges funding through the German Science Foun­dation (DFG) via SFB 755 project B08, SFB 937 project A13
and the Volkswagen foundation within the Niedersachsen Israel
framework (MWK-VWZN2722). T S acknowledges funding
through SFB 755 project C10, SFB 937 project A11, and BMBF
project 05K16MG2.
ORCID iDs
Toshio Ando https://orcid.org/0000-0001-8819-154X
H Colin-York https://orcid.org/0000-0002-6585-3237
P J de Pablo https://orcid.org/0000-0003-2386-3186
Christian Eggeling https://orcid.org/0000-0002-3698-5599
Jacob P Hoogenboom
https://orcid.org/0000-0003-45398772
Kris P F Janssen https://orcid.org/0000-0002-2181-9635
Judith Klumpermann https://orcid.org/0000-0003-4835-6228
Nalan Liv https://orcid.org/0000-0003-2654-9117
Florian Rehfeldt https://orcid.org/0000-0001-9086-3835
Iwan A T Schaap https://orcid.org/0000-0001-6665-5104
References[1] de Boer P, Hoogenboom J P and Giepmans B N G 2015
Correlated light and electron microscopy: ultrastructure
lights up! Nat. Methods 12 503–13[2] Timmermans F J and Otto C 2015 Review of integrated lightand electron microscopy Rev. Sci. Instrum. 86 011501[3] Agronskaia A V, Valentijn J A, van Driel L F,
Schneijdenberg C T W M, Humbel B M, van Bergenen Henegouwen P M P, Verkleij A J, Koster A J andGerritsen H C 2008 Integrated fluorescence andtransmission electron microscopy J. Struct. Biol. 164 183–9[4] Karreman M A, Agronskaia A V, van Donselaar E G,
Vocking K, Fereidouni F, Humbel B M, Verrips C T,
Verkleij A J and Gerritsen H C 2012 Optimizing immunolabeling for correlative fluorescence and electron
microscopy on a single specimen J. Struct. Biol. 180 382–6[5] Kukulski W, Schorb M, Welsch S, Picco A, Kaksonen M and
Briggs J A G 2011 Correlated fluorescence and 3D electron
microscopy with high sensitivity and spatial precision
J. Cell Biol. 192 111–9[6] Peddie C J, Blight K, Wilson E, Melia C, Marrison J, Carzaniga R,
Domart M C, O’Toole P, Larijani B and Collinson L M 2014
Correlative and integrated light and electron microscopy ofin-resin GFP fluorescence, used to localise diacylglycerol in
mammalian cells Ultramicroscopy 143 3–14
[7] Paez-Segala M G et al 2015 Fixation-resistant photoactivatable
fluorescent proteins for CLEM Nat. Methods 12 215–8[8] Faas F G A et al 2013 Localization of fluorescently labeled
structures in frozen-hydrated samples using integrated lightelectron microscopy J. Struct. Biol. 181 283–90[9] Iijima H, Fukuda Y, Arai Y, Terakawa S, Yamamoto N and
Nagayama K 2014 Hybrid fluorescence and electron cryomicroscopy for simultaneous electron and photon imaging
J. Struct. Biol. 185 107–15[10] Haring M T, Liv N, Zonnevylle A C, Narvaez A C,
Voortman L M, Kruit P and Hoogenboom J P 2017
Automated sub-5 nm image registration in integrated
correlative fluorescence and electron microscopy using
cathodoluminescence pointers Sci. Rep. 7 43621
38

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001[46] Sambur J B, Chen T-Y, Choudhary E, Chen G, Nissen E J,
Thomas E M, Zou N and Chen P 2016 Sub-particle reaction
and photocurrent mapping to optimize catalyst-modified
photoanodes Nature 530 77–80[47] Karreman M A, Buurmans I L C, Agronskaia A V, Geus J W,
Gerritsen H C and Weckhuysen B M 2013 Probing thedifferent life stages of a fluid catalytic cracking particle
with integrated laser and electron microscopy Chem. Eur. J.
19 3846–59[48] Van Loon J, Janssen K P F, Franklin T, Kubarev A V,
Steele J A, Debroye E, Breynaert E, Martens J A
and Roeffaers M B J 2017 Rationalizing acid zeolite
performance on the nanoscale by correlative fluorescenceand electron microscopy ACS Catal. 5234–42[49] Kennes K, Demaret C, Van Loon J, Kubarev A V, Fleury G,
Sliwa M, Delpoux O, Maury S, Harbuzaru B andRoeffaers M B J 2017 Assessing inter and intra-particle
heterogeneity in alumina-poor H-ZSM-5 zeolites
ChemCatChem 9 3440–5[50] Kubarev A V, Janssen K P F and Roeffaers M B J 2015
Noninvasive nanoscopy uncovers the impact of thehierarchical porous structure on the catalytic activity ofsingle dealuminated mordenite crystals ChemCatChem
7 3646–50[51] Yuan H et al 2016 Degradation of methylammonium lead
iodide perovskite structures through light and electron beam
driven ion migration J. Phys. Chem. Lett. 7 561–6[52] Yuan H, Debroye E, Caliandro G, Janssen K P F, vanLoon J, Kirschhock C E A, Martens J A, Hofkens J andRoeffaers M B J 2016 Photoluminescence blinking of
single-crystal methylammonium lead iodide perovskite
nanorods induced by surface traps ACS Omega 1 148–59[53] Manser J S, Christians J A and Kamat P V 2016 Intriguing
optoelectronic properties of metal halide perovskites Chem.
Rev. 116 12956–3008[54] Ersen O, Florea I, Hirlimann C and Pham-Huu C 2015
Exploring nanomaterials with 3D electron microscopy
Mater. Today 18 395–408[55] Debroye E, Van Loon J, Gu X, Franklin T, Hofkens J,
Janssen K P F and Roeffaers M B J 2016 Assessing
photocatalytic activity at the nanoscale using integrated
optical and electron microscopy Part. Part. Syst. Charact.
33 412–8[56] Debroye E, Van Loon J, Yuan H, Janssen K P F, Lou Z, Kim S,
Majima T and Roeffaers M B J 2017 Facet-dependent
photoreduction on single ZnO Crystals J. Phys. Chem. Lett.
8 340–6[57] de Jonge N and Ross F M 2011 Electron microscopy of
specimens in liquid Nat. Nanotechnol. 6 695–704[58] de Jonge N, Peckys D B, Kremers G J and Piston D W
2009 Electron microscopy of whole cells in liquid withnanometer resolution Proc. Natl Acad. Sci. 106 2159–64[59] de Jonge N 2018 Correlative light microscopy and liquidphase scanning transmission electron microscopy oflabelled membrane proteins in whole mammalian cells
J. Microsc. 269 134–42[60] Woehl T J, Kashyap S, Firlar E, Perez-Gonzalez T, Faivre D,
Trubitsyn D, Bazylinski D A and Prozorov T 2014
Correlative electron and fluorescence microscopy ofmagnetotactic bacteria in liquid: toward in vivo imaging Sci.
Rep. 4 6854
[61] Peckys D B, Korf U and de Jonge N 2015 Local variationsof HER2 dimerization in breast cancer cells discovered bycorrelative fluorescence and liquid electron microscopy Sci.
Adv. 1 e1500165[62] Shivanandan A, Deschout H, Scarselli M and Radenovic A
2014 Challenges in quantitative single molecule localization
microscopy FEBS Lett. 588 3595–602[30] Wolff G, Hagen C, Grünewald K and Kaufmann R 2016
Towards correlative super-resolution fluorescence andelectron cryo-microscopy Biol. Cell 108 245–58[31] Chang Y W, Chen S, Tocheva E I, Treuner-Lange A,
Löbach S, Søgaard-Andersen L and Jensen G J 2014
Correlated cryogenic photoactivated localization
microscopy and cryo-electron tomography Nat. Methods
11 737–9[32] Liu B et al 2015 Three-dimensional super-resolution protein
localization correlated with vitrified cellular context Sci.
Rep. 5 13017[33] Kim D, Deerinck T J, Sigal Y M, Babcock H P, Ellisman M H
and Zhuang X 2015 Correlative stochastic opticalreconstruction microscopy and electron microscopy PLoS
One 10 e0124581[34] Kopek B G et al 2017 Diverse protocols for correlative superresolution fluorescence imaging and electron microscopy ofchemically fixed samples Nat. Protocols 12 916–46[35] Collinson L and Verkade P 2015 Probing the future ofcorrelative microscopy J. Chem. Biol. 8 127–8[36] Ngo J T, Adams S R, Deerinck T J, Boassa D, RodriguezRivera F, Palida S F, Bertozzi C R, Ellisman M H andTsien R Y 2016 Click-EM for imaging metabolicallytagged nonprotein biomolecules Nat. Chem Biol. 12 459–65[37] Van Elsland D M, Bos E, de Boer W, Overkleeft H S,
Koster A J and van Kasteren S I 2016 Detection of
bioorthogenal groups by correlative light and electron
microscopy allows imaging of degraded bacteria inphagocytosis Chem. Sci. 7 752–8[38] Giepmans B N G, Deerinck T J, Smarr B L, Jones Y Z
and Ellisman M H 2005 Correlated light and electron
microscopic imaging of multiple endogenous proteins using
quantum dots Nat. Methods 2 743–9[39] Martell J D, Deerinck T J, Sancak Y, Poulos T L,
Mootha V K, Sosinsky G E, Ellisman M H and Ting A Y
2012 Engineered ascorbate peroxidase as a geneticallyencoded reporter for electron microscopy Nat. Biotechnol.
30 1143–8[40] Miles B T, Greenwood A B, Benito D, Tanner H, Galan M C,
Verkade P and Gersen H 2017 Direct evidence of lack ofcolocalisation of fluorescently labelled gold labels used incorrelative light electron microscopy Sci. Rep. 7 44666[41] Karreman M A, Mercier L, Schieber N L, Solecki G, Allio G,
Winkler F, Ruthensteiner B, Goetz J G and Schwab Y
2016 Fast and precise targeting of single tumor cells invivo by multimodal correlative microscopy J. Cell. Sci.
129 444–56[42] Janssen K P F, De Cremer G, Neely R K, Kubarev A V, Van
Loon J, Martens J A, De Vos D E, Roeffaers M B J and
Hofkens J 2014 Single molecule methods for the study ofcatalysis: from enzymes to heterogeneous catalysts Chem.
Soc. Rev. 43 990–1006[43] Merdasa A, Tian Y, Camacho R, Dobrovolsky A, Debroye E,
Unger E L, Hofkens J, Sundström V and Scheblykin I G
2017 ‘Supertrap’ at work: extremely efficient nonradiative
recombination channels in MAPbI3 perovskites revealed byluminescence super-resolution imaging and spectroscopy
ACS Nano 11 5391–404[44] De Cremer G, Roeffaers M B J, Bartholomeeusen E, Lin K,
Dedecker P, Pescarmona P P, Jacobs P A, De Vos D E,
Hofkens J and Sels B F 2010 High-resolution singleturnover mapping reveals intraparticle diffusion limitationin Ti-MCM-41-catalyzed epoxidation Angew. Chem., Int.
Ed. Engl. 49 908–11[45] Zhou X, Choudhary E, Andoy N M, Zou N and Chen P 2013
Scalable parallel screening of catalyst activity at the singleparticle level and subdiffraction resolution ACS Catal.
3 1448–53

39

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001[84] Eberle A L et al 2015 High-resolution, high-throughput
imaging with a multibeam scanning electron microscope
J. Microsc. 259 114–20[85] Lich B, Zhuge X, Potocek P, Boughorbel F and Mathisen C
2013 Bringing deconvolution algorithmic techniques to theelectron microscope Biophys. J. 104 500a[86] Mohammadi-Gheidari A and Kruit P 2011 Electron optics ofmulti-beam scanning electron microscope Nucl. Instrum.
Methods Phys. Res. A 645 60–7
[87] Kukulski W et al 2012 Precise, correlated fluorescencemicroscopy and electron tomography of lowicryl
sections using fluorescent fiducial markers Methods
Cell Biol. 111 235–57[88] Liv N et al 2013 Simultaneous correlative scanning electron
and high-NA fluorescence microscopy PLoS One
8 e55707[89] Peddie C J, Liv N, Hoogenboom J P and Collinson L M 2014
Integrated light and scanning electron microscopy of GFPexpressing cells Methods Cell Biol. 124 363–89[90] Iudin A, Korir P K, Salavert-Torres J, Kleywegt G J and
Patwardhan A 2016 EMPIAR: a public archive forraw electron microscopy image data Nat. Methods
13 387–8[91] Kaufmann R, Hagen C and Grünewald K 2014 Fluorescencecryo-microscopy: current challenges and prospects Curr.
Opin. Chem. Biol. 20 86–91[92] de Pablo P J, Schaap I A T, MacKintosh F C andSchmidt C F 2003 Deformation and collapse of
microtubules on the nanometer scale Phys. Rev. Lett.
91 098101[93] Ivanovska I L, de Pablo P J, Ibarra B, Sgalari G,
MacKintosh F C, Carrascosa J L, Schmidt C F and
Wuite G J L 2004 Bacteriophage capsids: tough nanoshells with complex elastic properties Proc. Natl Acad.
Sci. USA 101 7600–5[94] Pang H B, Hevroni L, Kol N, Eckert D M, Tsvitov M,
Kay M S and Rousso I 2013 Virion stiffness regulates
immature HIV-1 entry Retrovirology 10 4[95] Li S, Sieben C, Ludwig K, Höfer C T, Chiantia S,
Herrmann A, Eghiaian F and Schaap I A T 2014
pH-controlled two-step uncoating of influenza virus
Biophys. J. 106 1447–56[96] Greber U F 2016 Virus and host mechanics support
membrane penetration and cell entry J. Virol.
90 3802–5[97] Yeung T, Georges P C, Flanagan L A, Marg B, Ortiz M,
Funaki M, Zahir N, Ming W, Weaver V and Janmey P A
2005 Effects of substrate stiffness on cell morphology,
cytoskeletal structure, and adhesion Cell Motil.
Cytoskeleton 60 24–34[98] Sarkar A, Robertson R B and Fernandez J M 2004
Simultaneous atomic force microscope and fluorescencemeasurements of protein unfolding using a calibrated
evanescent wave Proc. Natl Acad. Sci. 101 12882–6[99] Ortega-Esteban A, Bodensiek K, San Martín C,
Suomalainen M, Greber U F, de Pablo P J and
Schaap I A T 2015 Fluorescence tracking of genome
release during the mechanical unpacking of single viruses
ACS Nano 9 10571–9[100] Pohl D 1986 IBM J. Res. Dev. 30 417–27[101] Gaiduk A, Kuhnemuth R, Antonik M and Seidel C A 2005
Optical characteristics of atomic force microscopy
tips for single- molecule fluorescence applications
ChemPhysChem 6 976–83[102] Thompson R E, Larson D R and Webb W W 2002 Precise
nanometer localization analysis for individual fluorescentprobes Biophys. J. 82 2775–83[103] Rust M J, Bates M and Zhuang X 2006 Sub-diffraction-limit
imaging by stochastic optical reconstruction microscopy(STORM) Nat. Methods 3 793–5[63] de Jonge N and Peckys D B 2016 Live cell electron
microscopy is probably impossible ACS Nano 10 9061–3[64] Bogner A, Thollet G, Basset D, Jouneau P H and Gauthier C
2005 Wet STEM: a new development in environmental
SEM for imaging nano-objects included in a liquid phase
Ultramicroscopy 104 290–301[65] Wojcik M, Hauser M, Li W, Moon S and Xu K 2015
Graphene-enabled electron microscopy and correlated
super-resolution microscopy of wet cells Nat. Commun.
6 1–6[66] Dahmke I N, Verch A, Hermannsdorfer J, Peckys D B,
Weatherup R S, Hofmann S and de Jonge N 2017 Graphene
liquid enclosure for single-molecule analysis of membraneproteins in whole cells using electron microscopy ACS
Nano 11 11108–17[67] Hirano K et al 2014 Electron microscopy of primary cell
cultures in solution and correlative optical microscopyusing ASEM Ultramicroscopy 143 52–66[68] Liv N, van Oosten Slingeland D S, Baudoin J P, Kruit P,
Piston D W and Hoogenboom J P 2016 Electron
microscopy of living cells during in situ fluorescencemicroscopy ACS Nano 10 265–73[69] Lucas M S, Günthert M, Gasser P, Lucas F and Wepf R 2012
Bridging microscopes: 3D correlative light and scanningelectron microscopy of complex biological structures
Methods Cell Biol. 111 325–56[70] Denk W and Horstmann H 2004 Serial block-face scanningelectron microscopy to reconstruct three-dimensional tissue
nanostructure PLoS Biol. 2 e329[71] Heymann J A W et al 2006 Site-specific 3D imaging of cells
and tissues with a dual beam microscope J. Struct. Biol.
155 63–73[72] Micheva K D and Smith S J 2007 Array tomography: a
new tool for imaging the molecular architecture andultrastructure of neural circuits Neuron 55 25–36[73] Bushby A J, Mariggi G, Armer H E J and Collinson L M 2012
Correlative light and volume electron microscopy: using
focused ion beam scanning electron microscopy to imagetransient events in model organisms Methods Cell Biol.
111 357–82[74] Oberti D, Kirschmann M A and Hahnloser R H R 2011
Projection neuron circuits resolved using correlative array
tomography Front. Neurosci. 5 1–8[75] Maco B et al 2013 Correlative in vivo 2 photon and focusedion beam scanning electron microscopy of cortical neurons
PLoS One 8 e57405[76] Collman X F et al 2015 Mapping synapses by conjugate lightelectron array tomography J. Neurosci. 35 5792–807[77] Blazquez-Llorca L et al 2015 Correlation of two-photon
in vivo imaging and FIB/SEM microscopy J. Microsc.
259 129–36[78] Beckwith M S et al 2015 Seeing a mycobacterium-infected
cell in nanoscale 3D: correlative imaging by lightmicroscopy and FIB/SEM tomography PLoS One 10 1–19
[79] Russell M R G et al 2017 3D correlative light and electron
microscopy of cultured cells using serial blockface scanningelectron microscopy J. Cell Sci. 130 278–91[80] Polishchuk E V, Polishchuk R S and Luini A 2013 Correlative
light–electron microscopy as a tool to study in vivo
dynamics and ultrastructure of intracellular structures
Methods Mol. Biol. 931 413–22[81] van Rijnsoever C, Oorschot V and Klumperman J 2008
Correlative light-electron microscopy (CLEM) combining
live-cell imaging and immunolabeling of ultrathin
cryosections Nat. Methods 5 973–80[82] Fermie J et al 2018 Single organelle dynamics linked to
3D structure by correlative live-cell and 3D electron
microscopy Traffic 19 354–69[83] Zheng Z et al 2018 A complete electron microscopy volume of
the brain of adult drosophila melanogaster Cell 174 730–43
40Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001[134] Honigmann A, Walter C, Erdmann F, Eggeling C and
Wagner R 2010 Characterization of horizontal lipid
bilayers as a model system to study lipid phase separation
Biophys. J. 98 2886–94[135] Weatherill E E and Wallace M I 2015 Combining singlemolecule imaging and single-channel electrophysiology J.
Mol. Biol. 427 146–57[136] Weiss S 1999 Fluorescence spectroscopy of single
biomolecules Science 283 1676–83[137] Groulx N, Juteau M and Blunck R 2010 Rapid topology
probing using fluorescence spectroscopy in planar
lipid bilayer: the pore-forming mechanism of the toxin
Cry1Aa of Bacillus thuringiensis J. Gen. Physiol.
136 497–513[138] Peters R 2003 Optical single transporter recording: transportkinetics in microarrays of membrane patches Annu. Rev.
Biophys. Biomol. Struct. 32 47–67[139] Montilla-Martinez M et al 2015 Distinct pores forperoxisomal import of PTS1 and PTS2 proteins Cell Rep.
13 2126–34[140] Mannuzzu L M, Moronne M M and Isacoff E Y 1996
Science 271 213–6[141] Cha A and Bezanilla F 1997 Neuron 19 1127–40[142] Zifarelli G and Kusch J 2015 Advanced Patch-Clamp
Analysis for Neuroscientists ed A Korngreen (Berlin:
Springer)[143] Kusch J and Zifarelli G 2014 Biophys. J. 106 1250–7[144] Lavis L D and Raines R T 2014 ACS Chem. Biol.
9 855–66[145] Kalstrup T and Blunck R 2013 Proc. Natl Acad. Sci.
110 8272–7[146] Richler E, Shigetomi E and Khakh B S 2011 J. Neurosci.
31 16716–30[147] Kusch J, Biskup C, Thon S, Schulz E, Nache V,
Zimmer T, Schwede F and Benndorf K 2010 Neuron
67 75–85[148] Sanderson M J, Smith I, Parker I and Bootman M D 2014
Fluorescence microscopy Cold Spring Harbor Protocols
2014 071795[149] Leblond F, Davis S C, Valdes P A and Pogue B W 2010
Pre-clinical whole-body fluorescence imaging: review
of instruments, methods and applications J. Photochem.
Photobiol. B 98 77–94[150] Roberts T P and Mikulis D 2007 Neuro MR: principles J.
Magn. Reson. Imaging 26 823–37[151] Jochimsen T H and Moller H E 2008 Increasing specificity
in functional magnetic resonance imaging by estimationof vessel size based on changes in blood oxygenation
Neuroimage 40 228–36[152] Le Bihan D, Urayama S-I, Aso T, Hanakawa T and
Fukuyama H 2006 Direct and fast detection of neuronal
activation in the human brain with diffusion MRI Proc.
Natl Acad. Sci. 103 8263–8[153] Miller K L et al 2007 Evidence for a vascular contribution
to diffusion FMRI at high b value Proc. Natl Acad. Sci.
104 20967–72[154] Yuste R, MacLean J, Vogelstein J and Paninski L 2011
Imaging action potentials with calcium indicators Cold
Spring Harbor Protocols 2011 985–9[155] Bai R et al 2015 Simultaneous calcium fluorescenceimaging and MR of ex vivo organotypic corticalcultures: a new test bed for functional MRI NMR
Biomed. 28 1726–38[156] Nemoto T 2014 Development of novel two-photon microscopy
for living brain and neuron Microscopy 63 i7–8
[157] Cui M et al 2017 A proof-of-concept study for developing
integrated two-photon microscopic and magnetic
resonance imaging modality at ultrahigh field of 16.4 tesla
Sci. Rep. 7 2733

[104] Adio S, Senyushkina T, Peske F, Fischer N, Wintermeyer W
and Rodnina M V 2015 Fluctuations between multiple
EF-G-induced chimeric tRNA states during translocationon the ribosome Nat. Commun. 6 7442
[105] Ando T, Uchihashi T and Scheuring S 2014 Chem. Rev.
114 3120–88[106] Sakiyama Y, Mazur A, Kapinos L E and Lim R Y H 2016
Nat. Nanotechnol. 11 719–23[107] Shibata M, Uchihashi T, Ando T and Yasuda R 2015 Sci. Rep.
5 8724[108] Kodera N, Yamamoto D, Ishikawa R and Ando T 2010
Nature 468 72–6
[109] Uchihashi T, Iino R, Ando T and Noji H 2011 Science
333 755–8[110] Chiaruttini N, Redondo-Morata L, Colom A, Humbert F,
Lenz M, Scheuring S and Roux A 2015 Cell 163 866–79[111] Uchihashi T, Watanabe H, Fukuda S, Shibata M and Ando T
2016 Ultramicroscopy 160 182–96[112] Zhou L, Cai M, Tong T and Wang H 2017 Sensors 17 938[113] Hecht B, Sick B, Wild U P, Deckert V and Zenobi R,
Martin O J F and Pohl D W 2000 J. Chem. Phys.
112 7761–74[114] Palomba S and Novotny L 2009 Nano Lett. 9 3801–4[115] Roca-Cusachs P, Conte V and Trepat X 2017 Quantifying
forces in cell biology Nat. Cell Biol. 19 742–51[116] Sabass B, Gardel M, Waterman C and Schwarz U 2008 High
resolution traction force microscopy based on experimental
and computational advances Biophys. J. 94 207–20[117] Tambe D T et al 2013 Monolayer stress microscopy:
limitations, artifacts, and accuracy of recovered
intercellular stresses PLoS One 8 e55172[118] Legant W R, Miller J S, Blakely B L, Cohen D M,
Genin G M and Chen C S 2010 Measurement of
mechanical tractions exerted by cells in three-dimensional
matrices Nat. Methods 7 969–71[119] Steinwachs J et al 2016 Three-dimensional force microscopyof cells in biopolymer networks Nat. Methods 13 171–6[120] Stout D A et al 2016 Mean deformation metrics forquantifying 3D cell–matrix interactions without requiring
information about matrix material properties Proc. Natl
Acad. Sci. 113 2898–903[121] Soine J R D, Brand C A, Stricker J, Oakes P W, Gardel M L
and Schwarz U S 2015 Model-based traction force
microscopy reveals differential tension in cellular actin
bundles PLoS Comput. Biol. 11 e1004076[122] Owen L M et al 2017 A cytoskeletal clutch mediates cellularforce transmission in a soft, 3D extracellular matrix
Mol. Biol. Cell[123] Bergert M et al 2016 Confocal reference free traction force
microscopy Nat. Commun. 7 12814[124] Labernadie A et al 2014 Protrusion force microscopy reveals
oscillatory force generation and mechanosensing activityof human macrophage podosomes Nat. Commun. 5 5343
[125] Kronenberg N M et al 2017 Long-term imaging of cellularforces with high precision by elastic resonator interference
stress microscopy Nat. Cell Biol. 19 864–72[126] Colin-York H et al 2016 Nano Lett. 16 2633–8[127] Colin-York H et al 2017 Nat. Protocols 12 783–96[128] Fritzsche M et al 2016 Sci. Adv. 2 1–12
[129] Harke B et al 2008 Opt. Express 16 4154–62[130] Vlasov I I et al 2013 Nat. Nanotechnol. 9 54–8
[131] Bingen P et al 2011 Opt. Express 19 23716–26[132] Honigmann A et al 2012 A high resolution electro-optical
approach for investigating transition of soluble proteins tointegral membrane proteins probed by colicin A Biochem.
Biophys. Res. Commun. 427 385–91[133] Bartsch P, Walter C, Selenschik P, Honigmann A and
Wagner R 2012 Horizontal bilayer for electrical andoptical recordings Materials 5 2705–30
41

Topical Review

J. Phys. D: Appl. Phys. 51 (2018) 443001[163] Bernhardt M, Nicolas J D, Eckermann M, Eltzner B,
Rehfeldt F and Salditt T 2017 Anisotropic x-ray scatteringand orientation fields in cardiac tissue cells New J. Phys.
19 013012[164] Weinhausen B, Saldanha O, Wilke R N, Dammann C,
Priebe M, Brughammer M, Sprung M and Köster S
2014 Scanning x-ray nano-diffraction on living
eukaryotic cells in microfluidic environments Phys. Rev.
Lett. 112 088102[165] Priebe M, Bernhardt M, Blum C, Tarantola M,
Bodenschatz E and Salditt T 2014 Scanning x-ray
nanodiffraction on dictyostelium discoideum Biophys. J.
107 2662–73[166] Howells M R et al 2009 An assessment of theresolution limitation due to radiation-damage in x-ray
diffraction microscopy J. Electr. Spectr. Relat. Phenom.
170 4–12[158] Fratzl P, Jakob H F, Rinnerthaler S, Roschger P and
Klaushofer K 1997 Position-resolved small-angle x-ray
scattering of complex biological materials J. Appl.
Crystallogr. 30 765–9[159] Weinhausen B et al 2012 X-ray nano-diffraction on
cytoskeletal networks New J. Phys. 14 085013[160] Wilke R N, Priebe M, Bartels M, Giewekemeyer K, Diaz A,
Karvinen P and Salditt T 2012 Hard x-ray imaging
of bacterial cells: nano-diffraction and ptychographic
reconstruction Opt. Express 20 19232–54[161] Bernhardt M et al 2016 X-ray micro- and nanodiffraction
imaging on human mesenchymal stem cells anddifferentiated cells Biophys. J. 110 680–90[162] Eltzner B, Wollnik C, Gottschlich C, Huckemann S and
Rehfeldt F 2015 The filament sensor for near real-time
detection of cytoskeletal fiber structures PLoS One
10 e0126346

42