SIM0216

VOLUME 18 

MAY 2016 

69721 

2 

Official Partner of the EMS 

High-Throughput Confocal Imaging 

Integrated Raman-FIB-SEM 

Multi-Tip SPM 

Single Molecular Spectroscopy

Editorial 

Image: Alexander Tselev and Andrei Kolmakov, ORNL 

Nano-Imaging with Microwaves 

Martin Friedrich, 

Editor-in-Chief 

Thomas Matzelle, 

Scientific Editor 

Nanoscale imaging in liquids is quite 

challenging: High-resolution imaging 

techniques that use energetic electron 

beams and X-rays often have a destructive 

effect on the sample. 

Surface researchers at the Oak Ridge 

National Laboratory, Tennessee, and the 

National Institute of Standards and Technology, 

Gaithersburg, Maryland, have recently 

demonstrated a non-destructive 

method for imaging objects and processes 

on the nanoscale in a liquid environment. 

Alexander Tselev, Andrei Kolmakov 

and their colleagues recommend 

an “environmental chamber” to encapsulate 

the sample in a liquid. In detail, the 

chamber has a window made of an ultra-thin 

membrane, not thicker than 50 

nm. The amazing part of the methodological 

set-up is the tiny tip of a scanning 

probe microscope that moves across the 

membrane injecting microwaves into the 

chamber. A high-resolution map of the 

sample is revealed when recording the 

transmitted versus the impeded microwave 

signal. This new methodological approach 

of combining scanning probe microscopy 

with microwaves and ultrathin 

membranes is called scanning microwave 

impedance microscopy (sMIM). 

Neutron scattering and X-ray diffraction 

are typical methods of choice 

when imaging crystals and other highly 

oriented materials. A promising new 

method is now to hand for microscopists 

when studying less ordered materials, 

like living cells and processes such as ongoing 

chemical reactions. 

A microwave oven heats aqueous liquids, 

as we all know. However, the microwaves 

injected in this way in sMIM are 

millions of times weaker and they oscillate 

in opposite directions. Thus, sMIM 

only produces negligible heat and potentially 

destructive chemical reactions cannot 

proceed. “Our imaging is nondestructive 

and free from the damage frequently 

caused to samples, such as living cells or 

electro-chemical processes by imaging 

with X-ray or electron beams,” said first 

author of the original publication in ACS 

Nano, Alexander Tselev. “Its spatial resolution 

is better than that achievable with 

optical microscopes for similar in-liquid 

samples. The paradigm could become instrumental 

in gaining important insights 

into electro-chemical phenomena, living 

objects and other nanoscale systems existing 

in fluids.” 

The applicability of sMIM has already 

been proven on different samples, especially 

living cells. The ORNL-NIST team 

has demonstrated, that to detect properties 

which distinguish healthy cells from 

sick ones: “If you have microwaves, you 

can go variably in depth and get a lot of 

information about the living, biological 

cell membrane itself - shape and properties 

that depend very much on the 

chemical composition and water content, 

which in turn depend on whether the cell 

is healthy or not,” Tselev said. 

Although current experiments have 

provided promising results regarding the 

applicability of the method for observations 

close to the surface, due to the nature 

of microwaves, sMIM is feasible for 

seeing deeper inside the sample. 

In future the researchers will try to 

further reduce the thickness of the membrane 

and to use probes and image-processing 

algorithms to improve the sensitivity 

of the system and the spatial 

resolution in depth. 

The stage is set for exciting and challenging 

advances in this field. 

References 

Dawn Levy: ORNL-NIST team explores nanoscale 

objects and processes with microwave microscopy, 

www.ornl.gov (2016) 

Alexander Tselev et al.: Seeing Through Walls at 

the Nanoscale: Microwave Microscopy of Enclosed 

Objects and Processes in Liquids, ACS Nano, 10 (3), 

pp 3562–3570 (2016) 

G.I.T. Imaging & Microscopy 2/2016 • 3


Contents 

69721 

VOLUME 18 

MAY 2016 

2 


EDITORIAL 3 

NEWSTICKER 6 

EVENT CALENDER 8 

ANNOUNCEMENT 

Bioimaging: from Cells to Molecules / EMBL Events 9 

High-Throughput Confocal Imaging 

Integrated Raman-FIB-SEM 

Multi-Tip SPM 

Single Molecular Spectroscopy 

European Microscopy Congress 2016 10 

SPAOM 2016 11 

Webinar: Fluorescence Lifetime Imaging 12 

COVER 

High-Throughput Confocal 

Imaging of 3D Spheroids 

Cover images show examples of some of 

the many automated imaging applications 

that are enabled using the new ImageXpress 

Micro Confocal High-Content Imaging 

System from Molecular Devices, including 

3D-spheroid imaging, tissue imaging, slide 

scanning, protein co-localization, fast kinetic 

processes such as beating cardiomyocytes, 

and whole organism (e.g. zebrafish) imaging. 

With the ImageXpress Micro Confocal 

you can run 3D cellular assays with confocal 

results — at a speed you’d only expect from 

widefield screening. 

16 

READ & WIN 

Handbook of Fluorescence Spectroscopy and Imaging 13 

From Single Molecules to Ensembles 

RMS IN FOCUS 

mmc2017 – It’s Your Congress 14 

NEWS FROM EMS 

EMS Newsletter 53, May 2016 15 

COVER STORY 

High-Throughput Confocal Imaging of 3D Spheroids 16 

Screening Cancer Therapeutics 

O. Sirenko 

LIGHT MICROSCOPY 

PREVIEW: ISSUE 3 

coming out August 17, 2016 

Diffusion Measurements in C. Elegans Embryos 18 

Using Single Plane Illumination Microscopy Combined with Fluorescence 

Correlation Spectroscopy 

P. Struntz et al. 

Single Molecular Spectroscopy 21 

Parallel Lifetime and Imaging of Single Molecules 

A. Mantsch and A. Cadby 

Water Wetting on Sub-Micron Scale 

Leaf Surfaces Studied with In Situ Electron 

Microscopy 

M. Koch 

TEM Imaging and TKD Mapping 

Interaction of Nanoparticles Incorporated in a 

Nickel Matrix 

D. Dietrich, T. T. Lampke, A. A. Sadeghi 

4 • G.I.T. Imaging & Microscopy 2/2016

Quality Control of Fluorescence Imaging Systems 24 

A New Tool for Performance Assessment and Monitoring 

A. Royon and N. Converset 

Observing the 3rd Dimension 28 

A Simple Way to Upgrade Common Microscopes for Sample Rotation 

T. Bruns et al. 

 

 

 

SCANNING PROBE MICROSCOPY 

The Multimeter at the Nanoscale 31 

Charge Transport at the Nanoscale Measured by a Multi-Tip 

Scanning Probe Microscope 

B. Voigtländer 

ELECTRON MICROSCOPY 

Integrated Raman – FIB – SEM 34 

A Correlative Light and Electron Microscopy Study 

F. Timmermans et al. 

 

Spectra of Electrons Emerging from PMMA 38 

Monte Carlo Simulation of Electron Energy Distributions 

M. Dapor 

Stemming Unwanted Interference 40 

Resolution Improvement by Incoherent Imaging with ISTEM 

F. Krause 

 

 

Electro-Optical Characterization of 3D-LEDs 44 

Nondestructive Inspection of 4” Wafers in Bird’s Eye View by an FE-SEM 

J. Ledig et al. 

PRODUCTS 47 

INDEX / IMPRINT 

INSIDE BACK COVER 

 

 

Congratulation 

The winner of Read & Win issue 

1/2016 is Pawel Drozdzal from 

Adam Mickiewicz University 

Polen. 

The next prize draw is on page 50 

 


NEWSTICKER 

High-Resolution Microscopy 

New Open Source Software 

With their special microscopes, 

experimental physicists 

can already observe single 

molecules. However, 

unlike conventional light microscopes, 

the raw image 

data from some ultra-high 

© University of Bielefeld 

resolution instruments first 

have to be processed for an image to appear. For the ultra-high resolution fluorescence 

microscopy that is also employed in biophysical research at Bielefeld 

University, members of the Biomolecular Photonics Group have developed a 

new open source software solution that can process such raw data quickly and 

efficiently. 

Original publication: 

Marcel Müller et al.: Open source image reconstruction of super-resolution 

structured illumination microscopy data in ImageJ, Nature Communications 

(2016) doi:10.1038/ncomms10980 

Laser Technology 

Changing the Orbital Angular Momentum of Laser Beams 

Researchers from South Africa and Italy 

demonstrating a new type of laser that is 

able to produce laser beams ‘with a 

twist’ as its output. These so-called vector 

vortex beams are represented on a 

higher-order Poincare sphere. Using geometric 

phase inside lasers for the first 

time, the work opens the way to novel 

lasers for optical communication, laser 

© University of the Witwatersrand 

machining and medicine. 


Darryl Naidoo et al.: Controlled generation of higher-order Poincaré sphere 

beams from a laser, Nature Photonics (2016) doi: 10.1038/nphoton.2016.37 

More information: 

http://bit.ly/IM-22016-b 


: http://bit.ly/IM-22016-a 

CLAIRE 

Super-Resolution Imaging 

Multiplexed Morse Signals from Cells 

How many sorts, in how many copies? 

The biochemical processes that take 

place in cells require specific molecules 

to congregate and interact in specific locations. 

A novel type of high-resolution 

microscopy developed at the Max 

Planck Institute of Biochemistry in Martinsried, 

Germany and Harvard Univer- 

© MPI Biochemistry 

sity, USA, already allows researchers to visualize these molecular complexes and 

identify their constituents. Now they can also determine the numbers of each molecular 

species in these structures. Such quantitative information is valuable for 

the understanding of cellular mechanisms and how they are altered in disease 

states. 


Ralf Jungmann et al.: Quantitative super-resolution imaging with qPAINT, 

Nature Methods (2016) doi: 10.1038/nmeth.3804 

Non-Invasive Electron Microscopy for Soft Materials 

Using the Molecular 

Foundry’s imaging capabilities, 

scientists developed 

a technique, called 

“CLAIRE,” that allows 

the incredible resolution 

of electron microscopy 

© Molecular Foundry to be used for non-invasive 

imaging of biomolecules and other soft matter. The new technique offers 

both clarity and speed. CLAIRE could lead to the understanding of key biological 

processes and help accelerate the development of new technologies such as 

high-efficiency photovoltaic cells. 

Original publications: 

Connor G. Bischak et al.: Cathodoluminescence-activated nanoimaging: Noninvasive 

near-field optical microscopy in an electron microscope, Nano Letters 

(2015) doi: 10.1021/acs.nanolett.5b00716 


http://bit.ly/IM-22016-d 


http://bit.ly/IM-22016-f 

MOZART 

Imaging Cells and Tissues 

under the Skin 

Scientists have many tools at their disposal 

for looking at preserved tissue 

under a microscope in incredible detail, 

or peering into the living body at 

lower resolution. What they haven’t 

had is a way to do both: create a threedimensional 

real-time image of individual 

cells or even molecules in a living 

animal. Now, Stanford scientists 

have provided the first glimpse under 

the skin of a living animal, showing intricate 

real-time details in three dimensions 

of the lymph and blood vessels. 

The technique, called MOZART 

(for MOlecular imaging and characteri- 

Zation of tissue noninvasively At cellular 

ResoluTion), could one day allow 

scientists to detect tumors in the skin, 

colon or esophagus, or even to see the 

abnormal blood vessels that appear in 

the earliest stages of macular degeneration 

– a leading cause of blindness. 


http://bit.ly/IM-22016-g 


Newsticker 

Electron Microscopy 

Real-Time Direct Observation of Atom Movements 

Atomic motion in a crystalline 

oxide that was used as a cathode 

in Lithium-ion batteries was directly 

demonstrated by state-ofan-art 

transmission electron microscopy, 

revealing the transient 

pathway of a chemical ordering 

reaction. Researchers from Korea 

have successfully demonstrated 

© KAIST 

how the cation ordering occurs in 

Li(Mn 1.5 Ni 0.5 )O 4 spinel, which is a promising cathode material for high-voltage 

Li-ion batteries. 


Hyewon Ryoo et al. : Frenkel-Defect-Mediated Chemical Ordering Transition in a 

Li-Mn-Ni Spinel Oxide, Angewandte Chemie, (2015) doi: 10.1002/ 

ange.201502320 


http://bit.ly/IM-22016-e 

Medical Imaging 

Breaking Bonds for Probes and Drugs 

A chemical procedure developed 

by an all-RIKEN research team has 

the potential to enhance the usefulness 

of positron emission tomography 

(PET) for discovering 

new drugs and diagnosing diseases. 

Compounds known as 

© RIKEN 

fluoroarenes are suitable starting materials for making fluorine-containing 

probes. But to transform them into useful probes requires breaking one of the 

strongest bonds in nature the carbon–fluorine bond. This is the key process that 

researchers have now achieved. They used a nickel–copper catalyst to break the 

carbon–fluorine bond in a way that permits non-radioactive fluorine-19 atoms to 

be swapped with radioactive counterparts, fluorine-18 atoms. 


Niwa T. et al.: Ni/Cu-catalyzed defluoroborylation of fluoroarenes for diverse 

C–F bond functionalizations, Journal of the American Chemical Society (2015) 

doi: 10.1021/jacs.5b10119 


http://bit.ly/IM-22016-h 

Look Sharp! 

PRECISION POSITIONING SOLUTIONS FOR MICROSCOPY 

Low-profi le XY stage 

with piezomotor drives 

Piezo tip/tilt mirrors 

for laser scanning 

Piezo XY and Z positioner 

for scanning, tracking and 

focusing 

PIFOC ® objective scanners 

with nanometer precision 

and travel up to 2 mm 

PI – The Broadest and Deepest Portfolio in Precision Motion Technologies. 

Physik Instrumente (PI) GmbH & Co. KG · www.pi.ws · info@pi.ws · +49 721 4846-0 

MOTION | POSITIONING 


EVENT CALENDAR 

APRIL HIGHLIGHT 

Ultrapath XVII 

11-15 July 2016 

Lisbon, Portugal 

http://congress.ultrapathxviii.org 

© Tupungato - Fotolia.com 

Microscopy & Microanalysis 

24-28 July 2016 

Columbus, Ohio, USA 

www.microscopy.org/MandM/2016 

MAY HIGHLIGHT 

AUGUST HIGHLIGHT 

19th International Conference 

on Non-Contact Atomic Force Microscopy 

25-29 July 2016 

Nottingham, UK 

http://ncafm2016.iopconfs.org/ 

© styxclick - Fotolia.com 

More Events on 

www.imaging-git.com/events 

© Lucian H Milasan - Fotolia.com 

2016 

Inter/Micro 6-10 June Chicago, USA www.mcri.org/v/101/InterMicro 

GerBI Core Facility Management Course 6-10 June Konstanz, Germany www.germanbioimaging.org/wiki/index.php/FMC_2016 

ICON Europe: International Conference 

on Nanoscopy 

7-10 June Basel, Switzerland www.icon-europe.org 

SCANDEM: Annual Conference of the Nordic 

Microscopy Society 

7-10 June Trondheim, Norway www.ntnu.edu/physics/scandem2016 

1st International Conference on Helium Ion 

Microscopy and Emerging Focused Ion Beam 8-10 June Luxembourg City http://hefib2016.list.lu/ 

Technologies 

Introducing Bioimaging: From Cells to Molecules 14-15 June Cambridge, UK http://selectbiosciences.com/conferences/index.aspx?conf=BC2016 

15th International Congress of Histochemistry 

and Cytochemistry 

19-22 June Istanbul, Turkey www.ichc2016.com 

Ultrapath XVII 11-15 July Lisbon, Portugal http://congress.ultrapathxviii.org 

GerBI Annual Community Meeting 11-13 July Fulda, Germany 

www.germanbioimaging.org/wiki/index.php/Annual_community_ 

meeting_2016 

International School on Fundamental Crystallography 

with applications to Electron Crystallography 

27 June-2 July Antwerp, Belgium 

www.uantwerpen.be/en/summer-schools/fundamental-electroncrystallography 

Microscopy & Microanalysis 24-28 July Columbus, USA www.microscopy.org/MandM/2016 

19st International Conference on 

Non-Contact Atomic Force Microscopy 

25-29 July Nottingham, UK http://ncafm2016.iopconfs.org 

16th European Microscopy Congress 28 Aug-2 Sep Lyon, France http://emc2016.fr/en/ 

Light Sheet Fluorescence Microscopy Conference 31 Aug-3 Sep Sheffield, UK www.rms.org.uk/discover-engage/event-calendar/lsfm2016.html 

Tomography for Scientific Advancement 

Symposium (ToScA) 

Spanish-Portuguese Meeting for Advanced 

Optical Microscopy 

6-7 September London, UK 

4-7 October Bilbao, Spain www.spaom2016.eu 

www.nhm.ac.uk/our-science/departments-and-staff/ 

core-research-labs/imaging-and-analysis-centre/tosca.html 


Introducing Bioimaging: From Cells to Molecules 

Cambridge, UK, June 14-15, 2016 

ANNOUNCEMENT 

This conference aims to address the challenges 

posed by modern imaging applications for life 

sciences and explore the benefits that can be 

achieved from doing so. 

If you utilize bioimaging techniques in 

your research or workflows, you will benefit 

from the expert knowledge of research 

leaders who are helping to define new 

parameters for experimentation and improve 

outcomes for those wishing to image 

cells, molecules and biological processes. 

Agenda Topics: 

▪▪ 

3D + Time Imaging 

▪▪ 

Correlative Imaging 

▪▪ 

Image Analysis 

▪▪ 

Probes & Biosensors 

▪▪ 

Single Molecule Imaging 

▪▪ 

Super-resolution Microscopy 

Speakers include, as Keynotes: 

▪▪ 

▪▪ 

Francesco Pavone, Principal Investigator, 

LENS, University of Florence 

Ralf Jungmann, Group Leader, Max 

Planck Institute of Biochemistry 

The agenda is available to view on the 

website. You can present your research 

on a poster while attending the meeting. 

Poster Submission Deadline: 07 June 

2016. Visit the website for submission information 

now! SelectBio is offering 3 for 

2 on all delegate passes at Bioimaging: 

From Cells to Molecules! 

Contact 

delegatesales@selectbio.com 

Phone: +44 (0) 1787 315110 

http://selectbiosciences.com/ 

Register now at: 

http://bit.ly/Bioimaging-UK 

Events @ EMBL in Heidelberg, Germany 2016 

Date Courses More information 

3 - 8 July EMBL Course: Advanced Fluorescence Imaging Technique www.embl.de/training/events/2016/MIC16-02/index.html 

25 - 30 July EMBL Course: Super-Resolution Microscopy www.embl.de/training/events/2016/MIC16-03/index.html 

28 Aug - 05 Sep EMBO Practical Course: 

Cryo-Electron Microscopy and 3D Image Processing 2016 

25 - 27 Sep EMBL–Wellcome Genome Campus Conference: 

Big Data in Biology and Health 

www.embl.de/training/events/2016/CRY16-01/index.html 

www.embl.de/training/events/2016/BIG16-01/index.html 

Come and see us at OPTATEC, 

June 7-9, Hall 3, Booth G48 

Vecteezy.com 

www.piezosystem.com 

Precision in Motion 

precise – fast – reliable 

individual piezoelectric solutions

Annoucement 

EMC 2016: The City of Lights! 

Lyon, France, August 28 – September 2, 2016 

The City of Lyon is not only a World Heritage 

Site, it is also known as the City of Lights. Like 

the yearly festival of lights, EMC2016 aims to 

be innovative, startling and rich in experience. 

Scientific Program Highlights 

With a very successful abstract submission, 

EMC2016 in Lyon promises to offer a 

high quality selection of communications. 

EMC 2016 will thus be one of the European 

milestones in Microscopy! Starting 

with 9 Pre–Congress Training Courses, 

on 25 and 26 August, the EMC 2016 scientific 

program will be dense and contain 

no less than 47 scientific sessions: 

▪▪ 

4 posters sessions 

▪▪ 

2 EMS Meetings: General Assembly 

and Council 

▪▪ 

2 Award Ceremonies: European Microscopy 

Award and a Micrograph 

Contest Award 

Spotlight on the Exhibition 

3700 sqm exhibition and more than 85 

exhibitors from Europe and beyond. EMC 

2016 will support innovation and start– 

ups by dedicating specific space to start– 

ups companies. The objective is to provide 

them an attractive showcase at a 

cheaper price. The exhibition will also 

keep space to welcome around 20 industry 

workshops. 

their first images in Lyon: “Workers leaving 

the Lumière Factory”. 

In order to echo the invention of cinema 

in Lyon, EMC 2016 aims at highlighting 

the world of imaging and its 

close links with microscopy. Moreover 

imaging will be honored not only through 

a micrograph competition but also with 

the development of videos on in situ microscopy 

and many more initiatives to be 

discovered onsite. 

Don’t forget to register and book your 

hotel from now on! 

▪▪ 

▪▪ 

▪▪ 

▪▪ 

▪▪ 

6 Special Scientific Workshops 

6 Plenary Lectures 

9 Life Sciences Sessions 

9 Materials Science Sessions 

9 Instrumentation & Method Sessions 

Luminous Ideas 

Lyon is the city of the Lumière brothers, 

the first filmmakers in history who made 


www.emc2016.fr/en 


Spanish-Portuguese Meeting for Advanced Optical Microscopy 

Bilbao, Spain, October 5-7, 2016 

ANNOUNCEMENT 

The Spanish-Portuguese Meeting for Advanced 

Optical Microscopy (SPAOM2016) cordially invites 

all researchers, facility managers and industry 

representatives interested in advanced 

bioimaging to its first meeting in Bilbao. 

SPAOM 2016 has its roots in the bi-annual 

meeting of the Spanish Network 

for Advanced Optical Microscopy (RE- 

MOA) and it is co-organized for the first 

time with the Portuguese Platform of 

BioImage. 

The conference aims at promoting the 

Spanish and Portuguese bioimaging scientific 

community. SPAOM 2016 also has 

an international scope: it will host talks 

by European researchers providing a 

meeting point with the broader international 

community. Participants will also 

have the opportunity to attend handson 

workshops on the latest microscope 

instrumentation. Deadline for abstract 

submission is 30 June 2016. 

Categories are: 

▪▪ 

Single-molecule imaging, optical super-resolution 

and CLEM 

▪▪ 

Light-sheet microscopy and in vivo 

imaging 

▪▪ 

Functional Imaging and Multi-spectral 

Microscopy (FRET, FLIM and FCS) 

▪▪ 

Optogenetics 

▪▪ 

Microscopy in Biophysics 

▪▪ 

Microscopy in Neurobiology 

▪▪ 

Core Facility Managing 

▪▪ 

Applications in the biosciences: cancer 

and biomedical imaging 

We warmly invite you to Bilbao to learn, 

exchange knowledge, and build a fruitful 

network within the bioimaging community 

in Spain and Portugal. 

More information on SPAOM 2016: 

www.spaom2016.eu 

fluorescence 

lifetime 

imaging 

frame 

rate up to 

90 fps 

unique 

resolution 

1008 x 1008 

pixels 

pco.flim camera system 

Fluorescence Lifetime Imaging 

application simplified 

operation 

frequency 

5 kHz - 

40 MHz 

www.pco.de 

www.pco-tech.com

Announcement 

Fluorescence Lifetime Imaging 

Using Fluorescent Decay Rates to Identify Individual Fluorophores 

Lifetime or decay rate relates to the phenomenon 

of fluorescence, the spontaneous 

emission of light that appears after 

the excitation of a sample. This process, 

photoluminescence, is widely utilized 

across many areas, for tagging cells and 

cell fragments in order to observe metabolic 

processes, for differentiating between 

reaction products or for the characterization 

of any parameters that 

induce a change in the fluorescence, for 

example oxygen partial pressure. 

Although the benefit of using fluorescence 

lifetime as an additional analytical 

parameter has been known for years, 

it has not been used on a broader scale 

with the exception of single point measuring 

devices, which are connected to 

scanning applications, and some image 

intensifier based systems. If the reason 

for this has been the complexity of the 

systems, we would like to recommend 

this webinar, which presents a new, less 

complex approach. 

In this webinar, Dr. Gerhard Holst, 

Leader of the R&D Department at PCO, 

will explain Fluorescence Lifetime Imaging 

in the frequency domain revealing 

all information on the background theory 

for its practical use. His presentation of a 

dedicated camera system will follow. First 

the differences between time domain and 

frequency domain fluorescence lifetime 

measurements will be explained and the 

special CMOS image sensor introduced. 

The camera system and its main features 

and limitations will then be described 

and the first experimental data (FRET 

and endogenous fluorescence), that have 

been obtained will be showcased. Finally 

some considerations about the application 

will be presented and the performance 

compared to alternative methods 

and then discussed. If your work involves 

measuring the luminescence lifetimes 

for FRET, calibration of optical chemical 

sensors or endogenous fluorescence differentiation, 

or if you are looking for dynamic 

changes in this parameter, this introduction 

to the new measuring system 

could be relevant to your application. 

Webinar on Fluorescence Lifetime Imaging on 

Thursday June 28th, 14:00 (CET) 

Using Fluorescent Decay Rates to 

Identify Individual Fluorophores 

What is fluorescent decay rate? What 

impact does it have on image data analysis? 

What are the requirements for using this 

method? How can the experiment be carried 

out most effectively? This webinar will be 

given by GIT Verlag‘s, A Wiley Brand 

journal “Imaging & Microscopy” 

and PCO. 

Contact 

Dr. Gerhard Holst 

Forschungsleiter PCO 

Kelheim, Germany 

gerhard.holst@pco.de 

Register free of charge: 

http://bit.ly/Webinar-PCO 


EAD & WIN 

Handbook of Fluorescence 

Spectroscopy and Imaging 

From Single Molecules to Ensembles 

Providing much-needed information 

on fluorescence 

spectroscopy and microscopy, 

this ready reference covers 

detection techniques, data 

registration, and the use of 

spectroscopic tools, as well as 

new techniques for improving 

the resolution of optical 

microscopy below the resolution 

gap. Starting with the basic 

principles, the book goes 

on to treat fluorophores and 

labeling, single-molecule fluorescence 

spectroscopy and 

enzymatics, as well as excited 

state energy transfer, and super-resolution 

fluorescence 

imaging. Examples show how 

each technique can help in 

obtaining detailed and refined 

information from individual 

molecular systems. 

Jörg Enderleins studied 

physics at the Mechnikov University 

in Odessa, Ukraine 

from 1981 until 1986, and 

defended his PhD thesis at 

Humboldt University in Berlin, 

Germany in 1991. He is 

now professor at the Georg- 

August University Göttingen, 

Germany and head of the research 

group “Single Spectroscopy 

and Imaging for 

Win the book! 

See the box on 

page 50 

Biophyisics and Compley Systems”. 

His main research 

topic is the development of 

new single-molecule spectroscopic 

techniques for biophysics 

applications. 

Johan Hofkens, born in 

Hoogstraten, Belgium, in 1966 

received his master in Chemistry 

from KU Leuven in 1988, 

which was followed by a PhD 

in Sciences from KULeuven 

in 1993. In 2005 he was appointed 

research professor at 

the KULeuven. His research 

interests are fast spectroscopy, 

single molecule spectroscopy 

and optics. 

Markus Sauer was born 

1965 in Pforzheim. He studied 

chemistry in Karlsruhe, Saarbrücken, 

and Heidelberg, and 

finished his PhD in Physical 

Chemistry at the University 

of Heidelberg in 1995 under 

the guidance of Prof. Jürgen 

Wolfrum. He is now professor 

at the University of Würzburg. 

His research interests 

cover the development 

of new electron transfer sensors 

and probes as well as 

new single-molecule sensitive 

fluorescence spectroscopic 

techniques. 

NEW 

X-Cite ® 120LEDBoost 

More Powerful Than Ever! 

LED illumination for 

fluorescence microscopy 

High power, broad-spectrum 

fluorescence excitation 

Instant ON/OFF and exceptional 

field uniformity 

More powerful than the original 

X-Cite 120LED 

Quiet, vibration-free thermal management 

Visit us at 

Contact X-Cite today to learn more! 

May 24–27th 

Debrecen, Hungary 

BOOTH SB7 


Sauer, M.; Hofkens, J. ; Enderlein, J. 

Handbook of Fluorescence Spectroscopy 

and Imaging. From Single Molecules to 

Ensembles 

Wiley-VCH, 2011 

Hardcover 

ISBN:3527316698, 

www.excelitas.com 

x-cite@excelitas.com 

2260 Argentia Road, Mississauga 

Ontario L5N 6H7 CANADA 


RMS IN FOCUS 

In 2017 the mmc scientific programme 

will be set by the participants. 

mmc2017 – It’s Your Congress 

The Microscience Microscopy Congress returns 

to Manchester at the beginning of July 2017, 

and as always, it has something new to offer. 

Dr Peter O’Toole and Professor Rik Brydson explain 

how, for the first time, the microscopy research 

community will define the final scientific 

programme. 

The Microscience Microscopy Congress 

2017 will open in Manchester on 3 rd July 

2017. It will be home to Europe’s largest 

event of the year dedicated to microscopy 

and imaging. The Microscience series 

is well-known for introducing new 

features and there is a noticeable one for 

2017; one that will have a significant effect 

on the content of the final scientific 

programme. 

“One of the big changes for this year 

is the call for papers,” explained Dr. Peter 

O’Toole, mmc2017 Life Sciences 

Chair. “For previous events the session 

titles have been set in advance and the 

call has been made. This has advantages 

in that it makes a very clear statement 

as to what will be covered within 

the conference. However, it also has its 

drawbacks in that it can force submitters 

to shoehorn their work into a par- 

ticular session. As an organizer it is not 

uncommon to receive papers that don’t 

quite fit your session but which together 

would make a really great session of 

their own. The new keyword system will 

be much more accommodating and it 

means that the final program will be the 

result of exactly what has been submitted. 

It’s very exciting.” 

The new system removes any barriers 

to submission and should attract abstracts 

from across the full breadth of 

microscopy. It also allows for a program 

to truly represent the current state of 

microscopy. 

“In the past, sessions may have been 

set up to a year in advance. This creates 

a snap-shot of ‘then’ rather than ‘now’,” 

said Professor Rik Brydson, mmc2017 

Physical Sciences Chair. “What we are 

saying is, if you are using a microscope 

in a new way, or developing a new technique, 

or have achieved results that are 

crying out to be shared, then you have a 

place at mmc. We are, in effect, giving the 

conference to those of you who are doing 

the exciting work – giving you the opportunity 

to present your work and to define 

the content of the event. It means that 

the final programme will be as ‘now’ as it 

is possible to get.” 

Representatives of the organizers, the 

RMS, attend events all over the world and 

are always on the look-out for new ideas 

that will improve the experience for delegates, 

exhibitors and day-visitors. Many 

of these ideas find their way quickly into 

RMS events. The Society is also open to 

suggestions from its members and from 

further afield. “We are proud of the mmc 

series, but we are always looking to improve 

it,” added Dr. O’Toole, who is also 

an Honorary Secretary of the RMS. “In 

particular we want to help people get to 

the event. We are a charity that exists to 

further microscopy and to support those 

working with microscopes. There is still 

more than a year to go until the doors of 

the congress open, so there is time for us 

to respond to new ideas and suggestions 

as to how we can make it easier for you 

to attend. We want everyone with a voice 

in microscopy to be with us in 2017 so 

my advice is go to the congress website, 

see what is already planned, and start 

thinking about how you can be a part of 

what promises to be a great event. And, if 

you have an idea to make it even better, 

get in contact with us and let’s talk.” 

Full details of mmc2017 can be found at: 

www.mmc-series.org.uk 


nEws from EMS 

Roger Wepf, EMS President 

EMS Newsletter 53 

May 2016 

Nick Schryvers, EMS Secretary 

Dear EMS member, 

For the present year, EMS has received 

around 36 applications for scholarships 

for attending EMC2016, the 16 th European 

Microscopy Congress, organized from August 

28 till September 2 in Lyon, France. 

26 of those have been selected to receive 

financial support from EMS to present 

their results and learn from the specialists 

in an international environment. 

A few weeks ago the jury of the EMS 

Outstanding Paper Award has come to a 

decision for the round of 2015. 17 high 

quality papers were nominated (nearly the 

same number as last year), and the following 

three were selected as award winners: 

▪▪ 

1. Instrumentation and Technique 

Development: “Quantum coherent 

optical phase modulation in an ultrafast 

transmission electron microscope”, 

Armin Feist, Katharina E. Echternkamp, 

Jakob Schauss, Sergey V. 

Yalunin, Sascha Schafer & Claus Ropers; 

Nature 521, 200-203 (2015) 

▪▪ 

2. Materials Sciences: “Imaging screw 

dislocations at atomic resolution by 

aberration-corrected electron optical 

sectioning”, Yang, H., Lozano, J. G., 

Pennycook, T. J., Jones, L., Hirsch, P.B., 

Nellist, P.D.; Nature Communications 6, 

7266 (2015) 

▪▪ 

3. Life Sciences: “Imaging G proteincoupled 

receptors while quantifying 

their ligand-binding free-energy 

landscape”, David Alsteens, Moritz 

Pfreundschuh, Cheng Zhang, Patrizia 

M Spoerri, Shaun R Coughlin, Brian 

K Kobilka & Daniel J Müller; Nature 

Methods 12, 845-851 (2015) 

We sincerely congratulate the authors of 

these winning papers who will receive 

their awards during the award ceremony 

on Thursday, September 1, at EMC2016. 

We also thank the nominators of all papers 

and look forward to a new round by 

next January 2017 for papers published 

in 2016. 

Next to the annual Outstanding Paper 

Award, this year we also select the 

winners of the fourth round of the prestigious 

quadrennial European Microscopy 

Award, this time sponsored by 

JEOL. The two winners will present a 

special lecture at the award ceremony 

of EMC2016. 

In a few weeks we will open a call for 

applications for the EMS Extension in 

2017, a term which indicates the strong 

involvement of and support from EMS. 

Deadline for these applications is June 

30, 2016. 

At its meeting in March, the Executive 

Board reviewed the 14 nominations for 

members of the Executive Board from 

September 2016 till September 2020. 

This new Board will be elected at the 

EMS General Assembly in Lyon and we 

invite all members to join this assembly 

on Thursday, September 1. More information 

will be provided in due course. 

Also at this March meeting, the Board 

reviewed the five final pre-bids for organizing 

EMC2020. At present, proposals 

for the following venues have been received: 

Basel (Switzerland), Catania (Italy), 

Copenhagen (Denmark), Ljubljana 

(Slovenia), Maastricht (The Netherlands). 

All chairs have received the notes and 

comments from the Board members 

and will prepare a final bid by coming 

May 31. They will present their bid during 

the EMS General Council meeting 

at EMC2016 in Lyon on Tuesday, August 

30, where the final choice will be made. 

Contact 

Prof. Dr. D. Schryvers, Ph.D. 

Electron Microscopy for Materials Science (EMAT) 

Department of Physics 

University of Antwerp, Belgium 

nick.schryvers@uantwerpen.be 


COVER STORY 

High-Throughput 

Confocal Imaging of 3D Spheroids 

Screening Cancer Therapeutics 

Oksana Sirenko 1 

Fig. 1: A workflow for testing spheroids in a high-throughput screening environment. A single spheroid 

can be grown in a 96 or 384-well plate, treated with compound, and stained with a cocktail of dyes that 

can be imaged without washing. Spheroids may also be fixed if desired. (Right) Transmitted light images 

of HCT116 cells were taken over the course of 63 hours using Timelapse acquisition on the ImageXpress 

Micro High-Content Screening System to show the formation of a spheroid (10X objective). 

There is a growing interest in using 3D spheroids 

to screen for potential cancer therapeutics 

as they are believed to mimic tumor behavior 

more effectively than 2D cell cultures. 

We discuss some of the challenges of developing 

robust spheroid assays and how they can 

be addressed, thus enabling rapid imaging and 

analysis of 3D spheroids in microplates. 

Introduction 

In recent years there has been significant 

progress in development of in vitro aggregates 

of tumor cells for use as models for 

in vivo tissue environments. When seeded 

into a well of a low-attachment round 

bottom microplate, these aggregates will 

form a discrete spheroid. Spheroids are 

believed to mimic tumor behavior more 

effectively than regular two dimensional 

(2D) cell cultures because, much like tumors, 

they contain both surface-exposed 

and deeply buried cells, proliferating and 

non-proliferating cells, and a hypoxic 

center with a well-oxygenated outer layer 

of cells. Such 3D spheroid models are being 

successfully used in screening environments 

for identifying potential cancer 

therapeutics. Here we discuss some of the 

challenges of developing robust spheroid 

assays, and how they can be overcome. In 

particular we will focus on: 

▪▪ 

▪▪ 

Locating and focusing on the spheroid 

in every well so it can be imaged in a 

single field-of-view 

Optimizing the compound and staining 

treatment to ensure dye penetration 

and avoid disturbing the spheroid 

placement 

▪▪ 

Acquiring representative images 

throughout the 3D structure, minimizing 

out-of-focus or background signal 

from above and below the imaging 

plane 

▪▪ 

Rapidly analyzing the images to yield 

meaningful results from which conclusions 

can be drawn 

Spheroid Formation and Treatment 

We used the following method to 

form spheroids from cancer cell lines 

HCT116, DU145, and HepG2. Cells were 

cultured in flasks at 37 °C and 5% CO 2 

before detaching and seeding into 96 

or 384-well black plates with clear bottom 

U-shaped wells (Corning 4520 and 

3830, respectively) at densities of 1000- 

1500 cells/well in the appropriate media 

supplemented with fetal bovine serum 

(FBS). Within 24 h, a single spheroid 

formed in the bottom of each well and 

continued growing in size until it was 

used for experimentation after 2-4 days 

Fig. 2: (Top) Best focus projection of 15 images of 

an HCT116 spheroid taken with widefield optics. 

Software segmentation counted 817 nuclei. 

Nuclei were missed due to distortion by unfocused 

fluorescence on the edges of the spheroid 

and poorly detected dim cells in the center. (Bottom) 

Best focus projection of 15 images of an 

HCT116 spheroid taken with confocal optics. A 

more accurate number, 1078 nuclei, was counted. 

at 37 °C and 5% CO 2 (fig. 1). Spheroids 

may be cultured longer but the increasing 

size may impede stain penetration 

and imaging of the center-most cells. 

Here we describe assays used to determine 

the effects of the anti-cancer compounds: 

etoposide, paclitaxel, and Mitomycin 

C. Spheroid treatment began 

by adding compounds into the wells at 

10x concentration then incubating for 

1-4 days, depending on the mechanism 

being studied. Shorter durations were 

used to study apoptosis and longer durations 

for multi-parameter cytotoxicity 

studies. For drug treatments longer 

than 2 days, compound was refreshed 

every 2 days at a 1x concentration. 

Staining and Imaging Spheroids 

The examples shown here are from the 

development of an HCT116 spheroid assay 

for evaluating spheroid morphological 

changes in addition to the incidence of 

apoptotic cells in the well. After the compound 

treatment was completed, stains 

were combined into a single cocktail at 


COVER STORY 

Fig. 3: (Top) Montage of image thumbnails of HCT116 spheroids in a 96 well 

plate treated with compounds and imaged with a 10X Plan Fluor objective. 

Hoechst stained nuclei (blue) are overlaid with CellEvent Caspase 3/7 apoptosis 

marker (green). Untreated controls are in column 4 and a Caspase 3/7 

response is evident in columns 5–7 where Paclitaxel was serially diluted 1:3 

from 1 µM in Row A (replicates of 3 across). (Left) Eleven Z planes were 

combined into a 2D Maximum Projection image and analyzed with a simple 

custom module. Raw images showing low and high degree of apoptosis 

with their corresponding segmentation masks are shown (royal blue = 

nuclei, pink = apoptotic cells). (Right) By normalizing the amount of apoptosis 

as compared to untreated spheroids and plotting on a graph, it can be 

seen that Paclitaxel (green line) induces apoptosis at a much lower concentration 

than either Mitomycin C or Etoposide. 

Fig. 4: Toxic effect of Antimycin A on mitochondria. (Top) Overlay of Hoechst 

(blue) and MitoTracker (orange) images of spheroids treated with Antimycin 

A in increasing concentrations of 1, 22, 67, and 200 nM. (Bottom) Plotted 

average intensity values of mitochondria identified within the spheroid 

illustrate the effect of the drug. 

4-6x concentration and added 

directly to the media in the 

wells. Stains that require no 

washing were chosen to avoid 

disturbing the spheroids. 

Spheroids were visualized 

using the ImageXpress Micro 

High-Content Screening 

System (Molecular Devices) 

at either 10x or 20x magnification. 

In order to analyze 

responses of cells throughout 

the 3D structure, images 

were collected from different 

sity in the stack to generate 

the projection. Confocal optics 

provide the ability to image a 

thinner optical section of the 

spheroid than widefield optics. 

This significantly reduces 

the amount of background 

haze produced by fluorescence-emitting 

objects above 

and below the plane being acquired. 

It also generally allows 

better resolution of fine 

detail either at the subcellular 

level or between cells that 

are clustered or stacked upon 

each other as they are within 

a 3D structure. More accurate 

segmentation is often possible 

using a confocal image. 

In repeated experiments with 

spheroids, segmentation of 

nuclei from widefield images 

yielded counts ~20% lower 

than nuclei counted in confocal 

images (fig. 2). 

Screening Anti-Cancer Drugs 

with an Apoptosis Assay 

One class of anti-cancer drugs 

targets the extrinsic pathway 

of apoptosis to trigger cell 

death. To demonstrate an assay 

for apoptosis, HCT116 

spheroids cultured in 96 well 

plates for 3 days were treated 

with a dilution series of 4 different 

anti-cancer compounds 

for 24-48 h. After the compound 

treatment was completed, 

apoptosis was detected 

using both CellEvent Caspase 

and MitoTracker Orange reagents 

from Life Technologies. 

A 4X cocktail of the combined 

stains, including Hoechst nuclear 

stain, was added to the 

media in the wells. Stains that 

require no washing out were 

chosen to avoid disturbing the 

spheroids (fig. 3). 

Multiplexing a Mitochondria 

Membrane Potential Assay 

in the Screen 

In the apoptosis screen above, 

mitochondrial membrane potential 

can also be evaluated 

by adding MitoTracker Orange 

to the dye cocktail. Alternatively, 

drugs that inhibit 

tumor growth by affecting mitochondria 

metabolism can 

depths within the body of the 

spheroid to create a “stack” of 

images. That stack of images 

was then combined or “collapsed” 

into a single 2D projection 

image using a mathematical 

algorithm. In this case 

a collapsed image was generated 

using the Maximum 

Projection algorithm in the 

MetaXpress High-Content Image 

Acquisition and Analysis 

Software. This keeps the pixels 

with the brightest intenbe 

studied separately. The following 

demonstrates an assay 

using Antimycin A, a potent 

disruptor of mitochondrial 

membrane potential. After 4 

h treatment, mitochondria 

health was detectable based 

on the intensity of MitoTracker 

Orange within the spheroid 

cells. The MitoTracker either 

did not penetrate completely 

to the center of the large spheroids 

or the cells in the center 

do not have healthy mitochondria 

as noted by the interior 

appearing generally dimmer 

in the Mitochondria wavelength 

in the images (fig. 4). 

Rapidly Screen 3D 

Spheroids in Microplates 

The ability of in vivo 3D culture 

systems to produce human 

cancer cell spheroids of 

uniform size and the ability 

to screen spheroid response 

to treatment using automated 

high-throughput, high-content 

imaging is a significant step in 

facilitating more relevant testing 

of chemotherapeutic drug 

candidates. The ImageXpress 

Micro High-Content Confocal 

Imaging System and MetaXpress 

Image Analysis software 

allow rapid imaging and analysis 

of 3D spheroids in microplates 

for monitoring induced 

apoptosis and mitochondrial 

toxicity of anti-cancer drugs. 

For further information on 

optimizing acquisition parameters 

in spheroid screening assays, 

please refer to: Sirenko, 

O. et al., High-Content Assays 

for Characterizing the Viability 

and Morphology of 3D Cancer 

Spheroid Cultures. Assay 

and Drug Development Technologies, 

2015. 13 (7): 402-14. 

Affiliation 

1 

Molecular Devices, 

Sunnyvale, CA, USA 

Contact 

Grischa Chandy 

Sr. Product Marketing Manager 

grischa.chandy@moldev.com 

Sarah Piper 

Marketing Manager Europe 

sarah.piper@moldev.com 

Molecular Devices 

www.moleculardevices.com 



Diffusion Measurements in C. Elegans Embryos 

Using Single Plane Illumination Microscopy Combined with Fluorescence Correlation Spectroscopy 

Philipp Struntz 1 , Matthias Weiss 1 , Benjamin Eggart 2 

We have used a combination of single plane illumination 

microscopy (SPIM) and fluorescence 

correlation spectroscopy (SPIM-FCS) to quantify 

protein diffusion in zygotes of the nematode 

Caenorhabditis elegans. 

Introduction 

In order to understand biological processes 

it is essential to quantify the diffusion 

behavior of proteins in the spatially 

inhomogeneous environment of a living 

specimen. 

A well-established technique for local 

diffusion measurement is fluorescence 

correlation spectroscopy (FCS). 

By correlating the intensity fluctuations 

of the fluorescence (GFP) in a 

small focal spot it is possible to derive 

the diffusion behavior of labeled particles. 

However, in many cases one would 

like to carry out multiplexed data acquisition 

in order to obtain diffusion 

maps that assess diffusional transport 

throughout an inhomogeneous en- 

vironment. Therefore, image based 

FCS-techniques have been developed. 

For imaging dynamic processes in cells 

and multicellular systems SPIM combines 

rapid widefield detection with optical 

sectioning by detecting the fluorescence 

emission (GFP) of perpendicularly 

illuminated slices of a sample. Imaging 

only the illuminated slice results in reduced 

bleaching and allows for longterm, 

three-dimensional in vivo imaging 

at a high spatiotemporal resolution 

[1,3,4,9] with reduced background 

signals. 

In SPIM-FCS [5, 6] each pixel of an acquired 

image represents a measurement 

point for the diffusion behavior while the 

confined illumination by a thin sheet of 

light restricts the axial extension of the 

focal volume. In order to resolve the decay 

of the autocorrelation in each pixel’s 

intensity trace thousands of images 

have to be acquired at a very high frame 

rate (1000 to 25000 fps). New scientific 

complementary metal oxide semiconductor 

(sCMOS) cameras make it possible 

to resolve even the rapid diffusion of 

proteins in the cytoplasm of living cells 

without destroying the sample during 

measurement. 

Experiment 

For SPIM-FCS, we have used a modified 

version of our previously published SPIM 

setup [1] as depicted in figure 1a. The 

widened beam of a DPSS-laser (491.5 

nm) was focused in one dimension by a 

cylindrical lens on the back aperture of 

an objective to obtain the illumination 

light sheet. To achieve the small observation 

volumes needed for FCS measurements, 

we reduced the thickness of the 

light sheet to a waist FWHM of 1.2 ± 0.1 

µm in a small rectangular region. Suitable 

eggs from transgenic worm lines expressing 

GFP-tagged PLC1δ1 were extracted 

and positioned in the waist of the 

light sheet. By imaging the light sheet 

waist at the middle of the sCMOS camera 

(ORCA-Flash 4.0, Hamamatsu Photonics, 

Japan) we reduced the number of horizontal 

lines to be read out (fig.1d). In 

this way, frame rates of 1000 to 25000 

fps were possible. In the rolling shutter 



Fig. 1: a) Sketch of our SPIM-FCS-Setup: The cylindrical lens focuses the beam in the x-dimension onto 

the excitation objective to form a light sheet. Fluorescence (GFP) is collected perpendicular with a 

second objective and focused onto the sCMOS camera. b) Side-view of the objectives showing the light 

sheet in the focus. c) The light sheet was positioned in the upper half of the ellipsoidal embryo. d) 

Illustration of the small rectangular region of interest on the sCMOS sensor to achieve high framerates. 

mode of the sCMOS chip are two readout 

registers (one for each sensor half). 

After 9.7 µs two horizontal sensor lines 

with a width of 2048 pxls are read out. 

Reducing the number of horizontal pixels 

therefore increased the total acquisition 

speed. The emitted fluorescence signal 

was filtered by a single-band filter 

and collected by a tube lens (fig.1b). The 

setup was controlled via a custom-made 

Labview program using trigger signals to 

control the camera via the Hokawo imaging 

software (Hamamatsu Photonics 

Deutschland). For measurements in the 

cytoplasm of the embryo up to 20,000 

frames with exposure times in the range 

152 − 1004 µs were taken. We imaged a 

layer in the upper half of the egg in order 

to reduce scattering and aberrations in 

the acquisition (fig.1c). Although the sC- 

MOS sensor is very sensitive it was necessary 

to use fairly high excitation powers 

in the range of 0.8 – 20 mW (measured 

at the backaperture of the illuminationobjective) 

which exceeded typical powervalues 

used for gentle long-term imaging 

(∼ 100 μW, 50 ms exposure-time) to 

maintain reasonable signal-to-noise ratios 

(SNR ~2.8 at light levels of ~210 photons/4 

pixels [10]). The possibility of both 

on-chip (2x2 binning) and subsequent 

software-binning (3x3 binning) was used 

to further improve the SNR at the cost 

of spatial resolution. Because of the increased 

excitation power as compared to 

SPIM imaging, timetraces in each pixel 

had to be corrected for bleaching effects. 

The auto-correlation function (ACF) 

of the corrected time traces were calculated 

with an open-source data evaluation 

software (Quickfit 3.0 Beta, SVN: 

Märzhäuser TrayExpress. 

Automated sample handling for microscopy. 

www.marzhauser.com 

Improve Your Microscope. 



3891 [7]). ACF-curves were then fitted 

using a model for three-dimensional diffusion 

to extract diffusion coefficients. 

Further details are available in Refs [1,2] 

Results 

To test the performance of SPIM-FCS in 

a well-established model organism, we 

measured protein diffusion maps in the 

early embryo of C. elegans. The GFPtagged 

protein PLC1δ1 is a peripheral 

membrane protein with a large cytoplasmic 

pool. Previous studies [8] determined 

the cytoplasmic diffusion to be in 

the range of 8.1±2.0 μm 2 /s. The fast diffusion 

pushed the SPIM-FCS application 

to its limit. Figure 2a illustrates the intensity 

image of an acquired layer in the 

early embryo in the one-cell stage. The 

lower intensity in the cytoplasm compared 

to the high signal on the membrane 

indicates a low amount of free 

proteins in contrast to elevated protein 

levels bound to PIP2 lipids on the plasma 

membrane. The autocorrelation function 

of a single pixel is shown in figure 

2c. The acquisition speed of the camera 

is sufficient to catch even the rapid ACF 

decay due to cytoplasmic diffusion at a 

reasonable accuracy. The resulting diffusion 

map is shown in figure 2b. Pixels 

not including cytoplasmic sites, having 

poor SNR, or showing measurement 

artifacts were masked. The diffusion 

maps in the embryo reflect a considerable 

cytoplasmic heterogeneity. The distribution 

of diffusion coefficients is depicted 

in figure 2d. Pixel values of the 

shown measurement have a broad distribution 

around a median value of 9.7 

μm2/s. This value is in a good agreement 

to previous reports [8]. The width of the 

distribution is the combination of the 

actual distribution of diffusive behavior 

throughout the embryo's cytoplasm 

and the fluctuating quality of the fitting-procedure 

for single pixels. A single 

SPIM-FCS measurement corresponds to 

hundreds of single point-FCS measurements. 

Therefore the multiplexed approach 

has the advantage of excellent 

statistics and reveals spatial differences 

in the diffusion behavior. To improve the 

data quality further a better SNR and 

shorter delay are crucial. 

Fig. 2: a) Fluorescence image of C. elegans embryo in one-cell stage expressing GFP-tagged PLC1δ1. b) 

The diffusion map of the embryo from (a) indicates the heterogeneous environment of the cytoplasm 

(additional 3x3 binning has been performed). c) Autocorrelation-function from the intensity time-trace 

of a single pixel. Fitting curve shown in red with residuals below. d) Distribution of all diffusion coefficient 

values from the measurement shown in (b). 

Conclusion 

The performance of our custom-built 

SPIM-FCS setup and an sCMOS camera 

is demonstrated by measuring on the established 

model organism C. elegans. 

By reading out only a small region parallel 

to the center-line of the camera 

sensor extremely high framerates were 

achieved. This enabled the autocorrelation 

of fast fluctuations in the fluorescence 

intensity signal caused by molecular 

diffusion. Due to the good SNR 

even at this short exposure times the 

resulting autocorrelation functions revealed 

diffusion maps of a model protein 

inside the cytoplasm of early C. elegans 

embryos. The presented data is in 

agreement with previous reports [8] on 

the same protein construct using scanning 

FCS measurements. The demonstrated 

SPIM-FCS method is therefore 

well suited to uncover vital processes 

in developmental biology. More detailed 

information can be found in reference 

[2] and online. 

References 

All references and a longer version of 

this article are available online: 

http://bit.ly/IM-Eggart2 

Acknowledgements 

Financial support from the DFG (grant 

WE4335/3-1) is gratefully acknowledged. 

Worm strains were provided by 

the CGC, which is funded by NIH Office 

of Research Infrastructure Programs 

(P40 OD010440). We would like to thank 

Malte Wachsmuth (EMBL Heidelberg) 

for valuable discussions on SPIM-FCS, 

and Jan Krieger and Joerg Langowski 

(DKFZ Heidelberg) for input on data 

evaluation. 

Affiliations 

1 

University of Bayreuth, Chair for Experimental 

Physics I, Bayreuth, Germany 

2 

Hamamatsu Photonics Deutschland 

GmbH, Herrsching am Ammersee, 

Germany 

Contact 

Dr. Benjamin Eggart, Application Engineer 

Hamamatsu Photonics Deutschland GmbH 

beggart@hamamatsu.de 

www.hamamatsu.de 

Prof. Dr. Matthias Weiss 

University of Bayreuth 

Chair for Experimental Physics I 

matthias.weiss@uni-bayreuth.de 

Read more about SPIM: 

http://bit.ly/IM-SPIM 

More information on C. elegans: 

http://bit.ly/IM-C-elegans 

[1] 

All references: 

http://bit.ly/IM-Eggart2 


liGHT MICROSCOPY 

Single Molecular Spectroscopy 

Parallel Lifetime and Imaging of Single Molecules 

Adrian Mantsch 1 and Ashley Cadby 1 

Typical image of single molecule fluorescence 

emission of a PCDTBT sample dissolved in 

Zeonex at a concentration of 5ng/l. 

Conventional microscopy and spectroscopy techniques can accurately analyze the properties of polymeric 

materials but incapable of distinguishing between properties in bulk and nanoscale. For this 

purpose, single molecular spectroscopy has been developed, a method that is able to circumvent 

these limitations. In this paper we present an automated experimental method capable of characterizing 

in real time a large number of individual molecules. 

Introduction 

Conventional microscopy and spectroscopy, 

as well as scanning probe techniques 

are capable of accurately characterizing 

polymeric films. But the 

intrinsic properties of the ensemble differ 

greatly from those of individual molecules, 

which are heavily influenced by 

the various degrees of disorder available 

to a single polymer chain. Therefore, the 

optical properties of thin film and that 

Fig. 1: Fluorescence spectra of PCDTBT in thin film (A) and at single molecule scale (B) both spectra 

were taken at room temperature. 

of a polymer’s will be radically different 

[1]. For example, figure 1 shows the optical 

properties in bulk and at nanoscale 

of a commercially applicable conjugated 

polymer. The thin film displays a relatively 

featureless spectrum, containing a 

broad dominant peak at 680 nm as well 

as several minor shoulders. At single 

molecule level, the same material shows 

four distinct fluorescence peaks, at lower 

wavelengths. 

To overcome some of the limitations 

associated with conventional microscopy 

and scanning probe microscopy, a 

new experimental method emerged in 

the early 1990’s: Single Molecular Spectroscopy 

(SMS), initially as a cryogenic 

method [1-3]. Figure 2 illustrates the basic 

operating principle of SMS. 

Conventional microscopes are capable 

of detecting single molecules but due to 

the small distance between single chains, 

below the diffraction limit, the system 

will be unable to resolve individual molecules 

and an ensemble measurement will 

be taken averaging the optical properties 

off all the molecules within the diffraction 

limited collection region. This limitation 

can be overcome by diluting the material 

of study to a level where the space 

between emitting molecules is higher 

than the device’s optical resolution. 

Single molecule fluorescence spectroscopy 

measurements require a large 



number of chromophores to be analyzed 

in order to get a statistically relevant 

data set. Due to the manual operation 

of the typical experimental set-ups, single 

molecule spectroscopy is a lengthy 

procedure and data acquisition is a very 

time consuming process. In order to automate 

and optimize the method, we 

are using a custom built optical microscope 

based on Zeiss optics. Automation 

was achieved by implementing a 

specific hardware configuration, working 

in conjunction with a series of software 

packages. In its current form, our 

experimental method allows the automatic 

acquisition of data in several 

modes of operation: fluorescence intensity 

acquisition; image capture, spectroscopy 

and photoluminesence (PL) lifetime 

measurements. 

Fig. 2: The operating principle of 

Single Molecular Spectroscopy. An 

extremely dilute sample (5 ng/ ml) 

sample is deposited on to a surface; 

the average separation between 

each molecule is greater than the 

diffraction limit. Improvements in 

camera technology in the 1990 and 

the large separation between 

molecules allows for the photo-luminesce 

from a single molecule to 

be resolved. The inset shows a 

typical photo-luminesce spectra. 

Sample Position 

An important function of the system is 

accurate control the samples position. 

This is achieved by means of a Zaber A- 

series microscope stage for coarse positioning, 

and a nPoint piezo stage for fine 

positioning. In this configuration, there 

are two distinct modes of operation: 

manual and automatic. In manual mode, 

the user can freely move the sample either 

in coarse steps or in finer sub-micron 

steps; this allows the user to select 

areas of interest within a sample. 

The second operating mode involves 

the automatic movement, important in 

data acquisition. For this purpose, the 

software accompanying the piezo stage 

(nP Control) is equipped with a raster 

scanning mode. The sample is moved on 

one of the horizontal axes (noted for convenience 

as X) in equal steps of pre-determined 

size. Between the steps, a controller 

(nP LC403 controller) will trigger a 

Princeton Instrument ProEM 512 EMCCD 

camera or a secondary capture device, for 

acquiring data. When motion on the X axis 

is complete the sample will be moved one 

step on the perpendicular axis and the cycle 

is repeated. The result will be a raster 

pattern on the film surface. The system is 

highly flexible with all parameters of the 

raster pattern being determined by the 

user. That includes the number of steps on 

the X axis, the number of lines in the pattern 

(steps on Y axis) as well as the dwell 

time between each step (used in data acquisition). 

If necessary the raster pattern 

can be extended on the vertical axis, automatically 

repeating the horizontal raster 

scan. This operating mode is useful for 

acquiring data in “slices”, to create a 3D 

scan of the analyzed sample. 

Fluorescence Intensity 

and Extended Lifetime 

For all measurements a laser is focused to 

a tight spot on to the sample. When measuring 

fluorescence intensity, the diffraction 

limited laser spot is imaged using the 

EMCCD camera at each point of the scanners 

raster pattern. Optical filters are 

used to remove the laser light allowing 

only PL to be detected on the camera. The 

intensity of the spot is integrated over the 

point-spread function of the microscope 

to build up a map of PL intensity. A typical 

intensity image is given in figure 3a. 

Extended lifetime measurements are 

performed by means of Avalanche Photo 

Diode (APD) detector, provided by Photonic 

Solutions. We use time-correlated 

single photon counting methods to measure 

PL lifetimes (TCSPC), method based 

on detecting individual photons of a periodic 

signal, measuring detection times 

and reconstructing the waveform from 

the time measurements. TCSPC is possible 

because the intensity of low level 

high repetition rate signals is usually so 

low that the probability of detecting more 

photons in a single signal period is insignificant. 

Upon detecting a photon, a detector 

pulse in the signal period is measured. 

When a large enough number of photons 

has been measured, their distribution 

over the signal period time builds up 

and the result is a distribution probability, 

in the shape of a waveform, of the optical 

pulse [4]. Again for each point on the 

scanners raster pattern a full PL lifetime 

curve is collected, this requires a longer 

delay time, on the order of 300 ms. A typical 

lifetime curve is given in figure 3b. 

Figure 3 shows an example of data acquired 

by the described module. The controlling 

(Becker & Hickl) SPCM software, 

together with the nPoint controller and 

piezo stage, will automatically scan the 

sample surface providing a fluorescence 

intensity map (fig. 3A), as a preliminary 

analysis, as well as lifetime data (fig. 3B) 

for various types of samples. 

Spectroscopy 

For spectroscopy measurements, our 

custom set-up is equipped with a ProEM 

512 electron-multiplying charged couple 

device camera and an Acton SP2500 

spectrometer, both provided by Princeton 

Instruments [5]. The spectrometer is 

composed of a slit, for minimizing collection 

on both horizontal axes as well as 

a multi-grating turret containing a mirror 

for conventional imaging and a series 

of two gratings (150 and 300 grooves/ 

Further information on 

microscopy of single molecules: 

http://bit.ly/IM-SMI 

Read more about automation 

in modern microscopy: 

http://bit.ly/IM-auto 

[1] 


http://bit.ly/IM-Mantsch 



Fig. 3: Example of lifetime data, 

acquired for PCDTBT single molecules. 

(A) fluorescence intensity map, 

(B) typical lifetime decay curve. 

mm respectively) for fluorescence 

spectroscopy measurements. 

Again for each point of 

the raster scan the camera is 

triggered to collect a PL spectrum 

from the spot excited by 

the laser. 

Conclusions 

We have optimized and successfully 

implemented a custom 

experimental set-up and 

method for single molecular 

spectroscopy. By using a specific 

hardware configuration 

and software packages, the 

system is capable of automatically 

collect data from a large 

number of emitters, in various 

modes of acquisition: fluorescence 

intensity, fluorescence 

spectroscopy, image capture 

or lifetime measurements. The 

set-up offers a high degree of 

flexibility, allowing the characterization 

of many types of 

samples from thin polymeric 

films biological samples or fluorescent 

beads. 

We have 

developed 

a new 

breed... 

References 

All references are available 

online: 

http://bit.ly/IM-Mantsch 

Affiliation 

1 

The University of Sheffield, 

Department of Physics and 

Astronomy, Sheffield, United 

Kingdom 

Contact 

Adrian Mantsch 

The University of Sheffield 

Department of Physics and Astronomy 

Sheffield, United Kingdom 

a.mantsch@sheffield.ac.uk 

The latest ground-breaking 

innovation in Microscopy. 

It’s more than just confocal. 

Find out more at 

andor.com/newbreed


Quality Control of 

Fluorescence Imaging Systems 

A New Tool for Performance Assessment and Monitoring 

Arnaud Royon 1 and Noël Converset 2 

We have developed a new tool for the assessment 

and monitoring of most of the performances of 

fluorescence microscopes. We believe it can advantageously 

be integrated in the quality control 

process of core facilities where a certain level of 

performance for the end users must be assured. 

Context 

Although performance evaluation and 

quality control of fluorescence microscopes 

is a topic that appeared more 

than fifteen years ago in academic laboratories 

[1] and national regulatory 

agencies [2], it is still topical as it was in 

the program of the Core Facility Satellite 

Meeting of the 15th international ELMI 

meeting in 2015. Due to the increasing 

complexity of the instrumentation used 

for confocal and high-end wide-field flu- 

orescence imaging microscopy, national 

metrology institutes [3], microscope 

manufacturers [4], and more recently 

core facilities [5] have gotten involved in 

identifying, manufacturing and/or testing 

different tools, both hardware and software, 

to assess the numerous aspects of 

fluorescence microscopes. 

On the one hand, for the core facilities, 

it has become obvious that quality 

control of fluorescence microscopes 

is important, as they provide a charged 

service to microscope end users. In this 

sense, they have to assure, up to a certain 

level, the performances of their 

microscopes. Quickly identifying and 

solving microscope issues is therefore 

essential in order to prevent the acquisition 

of corrupted data and to minimize 

the machine downtime. That is why core 

facilities usually spend tens of thousands 

euros per year for the maintenance of 

their systems. 

On the other hand, for the microscope 

manufacturers, maintenance is not as effective 

as it could be for two main reasons. 

First, in average, one intervention of 

the maintenance service over two is not 

justified, as it is based on a wrong (human 

misinterpretation) in situ diagnosis 

of the system while it performs correctly. 

Second, when the system is faulty, the 

identification and fixation of the problem 

can require several interventions. Knowing 

in advance what the microscope issue 

is allows optimizing the maintenance, if it 

is necessary. This would reduce the maintenance 

time and increase the technician 

availability for others systems/facilities. 

For both these actors, a win-win opportunity 

could arise if an evaluation and 

monitoring tool, accepted from both sides, 



and dark fields, DIC (Differential Interference 

Contrast) and phase contrast. 

Each fluorescent pattern is designed for 

one or several performance assessments. 

Non-exhaustively, the slide allows to 

assess and monitor the following characteristics 

of a fluorescence microscope 

(confocal, spinning-disk and wide-field): 

Evenness of illumination, distortion of 

the field of view, parcentrality, parfocality, 

optical axis determination, chromatic 

lateral shifts, co-localization issues, 

stitching performance, stage repositioning 

accuracy, intensity response of the 

system, spectral response of the system, 

lateral resolving power, objective issues, 

three-dimensional (3D) reconstruction 

precision, distances in XY and Z, and 

scanning performance. 

INTRODUCING THE 

UC Series 

TECHSPEC ® ULTRA 

COMPACT LENSES 

Superior Performance 

for Small Sensors 

Fig. 1: (A) The slide, containing numerous fluorescent 

patterns (B). 

would allow to assess quickly and simply 

most of the performances of a fluorescence 

microscope. Thus, the core facilities 

would get the assurance they provide the 

best possible service, so that the end users 

get reliable data, while the microscope 

manufacturers would reduce their maintenance 

intervention time and would improve 

their knowledge of the malfunction 

sources, so that they can correct them. Besides, 

after an installation or an intervention, 

both parties could validate the performances 

of a system no longer on the 

basis of a subjective image of a biological 

sample, but on objective and quantified 

parameters. This would be a huge step 

towards quality control of fluorescence 

microscopes. 

Having perceived the significance of 

this issue, we have worked together to develop 

a new tool, for the performance assessment 

and monitoring of fluorescence 

microscopes. This tool aims to: first, validate 

a system at a t 0 origin time (after an 

installation and/or a maintenance); second, 

monitor the performances of a system 

over time; and third, detect any malfunction 

of a system. 

Evaluation Slide 

The device, basically a slide, consists of a 

custom glass substrate, set on a stainless 

steel carrier (fig. 1A). The carrier features 

the same dimensions as a standard 

microscope slide. Different fluorescent 

patterns (fig. 1B) are embedded inside 

the glass, at a depth emulating the presence 

of a microscope cover-slip. These 

patterns also exhibit a contrast in bright 

Spectral Features 

The patterns exhibit the following fluorescence 

spectral features. Excitation: 

The excitation ranges from 300 up to 650 

nm. The excitation efficiency is maximal 

at around 340 nm and drops towards the 

red wavelengths. Emission: The emission 

is a continuum starting from slightly 

above the excitation wavelength up to 

800 nm. Lifetime: Using FLIM (Fluorescence 

Lifetime Imaging Microscopy), two 

main decay components of (0.25 ± 0.05) 

ns and (2.50 ± 0.50) ns have been measured. 

Photo-stability: The intensity of 

the patterns may decrease, but this decrease 

is transient. The fluorescence intensity 

recovers to its initial value after 

some time. The recovery time depends on 

the irradiation conditions (power density, 

wavelength, pixel size, exposure time). 

Performance Assessment Examples 

The tests that can be performed with the 

slide are too numerous to be described 

individually in the framework of the present 

paper. We will therefore limit ourselves 

to three examples, illustrating 

nevertheless the potential of this tool. 

Lateral Chromatic Shifts 

Because the patterns can be excited from 

the UV up to the red, lateral shifts between 

different channels can be measured. 

The 1 mm² matrix of rings (blue inset 

in fig. 1B) allows doing that, not only 

in the center of the field of view as one 

would do with a bead, but in its whole. 

Figures 2A-D depict the matrix of rings 

imaged with three different channels 

(DAPI, GFP, and Texas Red), and the superposition 

of these channels, respec- 

BORIS 

ECKER 

Imaging 

Solutions 

Engineer 

Designed by Imaging Experts 

Manufacured by Perfectionists 

and Attractively Priced for You. 

CONTACT OUR IMAGING 

EXPERTS TODAY! 

2016 

EDUCATIONAL 

AWARD 

OPTICAL 

INSPIRING INNOVATION 

NICHOLAS 

SISCHKA 

Vision 

Solutions 

Specialist 

Take your chance 

and apply by 

30 June 2016. 

Welcome To 

GREG 

HOLLOWS 

Director, 

Imaging 

Business Unit 

UK: +44 (0) 1904 788600 

GERMANY: +49 (0) 721 6273730 

FRANCE: +33 (0) 8 20 20 75 55 

sales@edmundoptics.eu 

www.edmundoptics.eu/uc-series


tively. Figures 2F and 2G present the intensity 

profiles for the three channels for 

two rings of the image, one at the bottom 

left and the other one at the top right, 

respectively. One can see that the lateral 

shift between the three channels is 

no more than 130 nm, less than the lateral 

resolution of the system (here about 

200 nm) for the present Plan-Apochromat 

63×/1.4 objective. Plan-Apochromat 

means that the lateral shift between four 

different colors (dark blue, blue, green 

and red) must be less than the system 

lateral resolution. The results shown in 

figure 2 are in accordance with the manufacturer 

specifications, for the three 

present channels. 

Fig. 2: Confocal images (Plan-Apochromat 63×/1.4 objective) of the matrix of rings for three different 

channels (DAPI, A; GFP, B; and Texas Red, C), and the superposition of these channels (D). (E) Inset: 

Zoom of one ring for the three channels, and their superposition. Lateral shift between the three 

channels for two rings of the image, one at the bottom left (F) and the other at the top right (G). 

Fig. 3: Confocal images (40×/1.3 objective, λexc=405nm, Δλem=410-605 nm) of the 2 µm step “crossing 

stairs” in good conditions (A), and with the DIC slider partially inserted (B). 3D reconstruction of 

the stairs (D). 

Objective Issues and 3D 

Reconstruction Precision 

An interesting aspect of this technology 

is its ability to induce patterns at different 

depths of different lengths. We have 

designed four 3D patterns (red inset in 

fig. 1B), consisting of rings at different 

depths with different steps (5, 2, 0.5 and 

0.2 µm) surrounded by four pillars, featuring 

two “crossing stairs”. On a single 

2D image, it is therefore possible to have 

access to the spreading of the light in 3D 

from the rings at different planes, and to 

get a clue on the out-of-focus issues. 

The “crossing stairs” with a 2 µm step 

has been imaged with the same objective, 

in good conditions (fig. 3A) and with 

a DIC slider partially inserted in the optical 

path in order to simulate a problem 

(fig. 3B). On the image acquired with the 

partially inserted DIC slider (fig. 3B), one 

can see that the light spread by the infocus 

rings looks correct, while the light 

spread by the out-of-focus rings does not 

have a circular symmetry, unlike in figure 

3A, evidencing the presence of an 

issue. This is a simple and fast way to 

check the optical quality of a system. Besides 

this particular case, one can also 

observe with this method if a microscope 

objective is damaged or if there is dust 

or oil on it. 

Such a pattern can also be used to 

evaluate the accuracy of its reconstruction 

in 3D, as it is illustrated in figure 3C. 

In this figure, we can clearly see that the 

reconstruction is accurate, and that the 

Z-distances are those expected. 

System Intensity Response 

This technology also enables to control 

the fluorescence intensity of the patterns, 

up to 16 well-discriminated intensity levels 

following a warranted linear evolution. 

The pattern consists in 16 squares 

having different intensities (green inset 

in fig. 1B). It can be used to characterize 

the intensity response of the system, in 



Schneider - 

Kreuznach 

Industrial 

Optical 

Filters 

Fig. 4: (A) Wide-field fluorescence image (40×/0.95 objective, GFP channel, 1 second exposure time) of 

the 16 squares having different intensities. (B) Evolution of the mean intensity of each square versus 

the square number. The linear regression curve has a Pearson coefficient better than 0.99, evidencing 

a good linear response of the actual camera. (C) Screen shot of the Daybook-Z interface. 

terms of linearity range, sensitivity, and 

limit of saturation. 

Figure 4A shows an image of these 16 

squares. The mean intensity of each 

square has been extracted and plotted 

on a graph versus the square number. 

The evolution of the intensity levels follows 

a linear trend, with a Pearson correlation 

coefficient better than 0.99, evidencing 

a good linear response of the 

actual camera (fig. 4B). This analysis has 

been achieved with Daybook-Z, the companion 

analysis software of the Argo-Z 

slide, in less than a minute. Besides the 

intensity response of the system, Daybook-Z 

also allows to extract from images 

of the suitable patterns the illumination 

homogeneity, the field distortion, 

the spatial co-localization, the lateral resolving 

power, the stage repositioning 

accuracy and the spectral response of 

the system (fig. 4C). 

For the first time to our knowledge, 

there exists a technology that allows to 

assess and to monitor most of the performances 

of fluorescence microscopes 

over the same time scale as their lifetime. 

The presented slide satisfies the requirements 

listed by the broad community 

of fluorescence microscopists and 

the National Metrology Institutes [3]. This 

is a huge step towards quality control of 

these instruments. Besides, because all 

the patterns are accurately positioned, 

it is possible to fully automatize the assessment 

process, first in the acquisition 

of the images, secondly in the analysis 

through dedicated algorithms, and 

thirdly in the edition of quality management 

documents, including data, graphs, 

reports, etc. 

References 

All references are available online: 

http://bit.ly/IM-Royon2 

Affiliations 

1 

Argolight SA,Talence, France 

2 

Carl Zeiss SAS, Marly-le-Roi, France 

• For automated 

production 

• For laser applications 

• As cover glass 

• To increase contrast 

• For metrology 

• Custom designed and 

manufactured 

Conclusion 

Contact 

Dr. Arnaud Royon 

Argolight SA 

Institut Optique d’Aquitaine 

Talence, France 

a.royon@argolight.com 

www.argolight.com 

Read more about Fluorescence 

Lifetime Imaging: 

http://bit.ly/IM-FLIM 

Recent advances in 

instrumentation: http:// 

bit.ly/IM-calibration 


http://bit.ly/ 

[1] IM-Royon2 

www.schneiderkreuznach.com


Observing the 3rd Dimension 

A Simple Way to Upgrade Common Microscopes for Sample Rotation 

Thomas Bruns 1 , Sarah Bruns 1 , Herbert Schneckenburger 1 

In microscopy, samples are usually located on 

glass slides or in specific dishes and may, therefore, 

only be observed from one direction. This 

is especially unfavorable for three-dimensional 

samples where larger or more complex specimens 

or structures are to be studied. For that reason 

we developed a modular device allowing longitudinal 

axial rotation of the specimen up to 360°, 

independent of the sample size. It can be easily 

adapted to a variety of common microscopes for 

gaining deeper insights into the sample. 

Fig. 1: Device for rotation of three-dimensional samples to be fixed on a positioning stage of a 

microscope. 

Sample rotation in microscopy is getting 

into the focus. Within the last years some 

efforts were made to vary the perspective 

of sample illumination and/or observation 

(see for example Bradl et al. [1], Staier et 

al. [2] and Heintzmann and Cremer [3] for 

single cells and nuclei and Huisken and 

Stainier [4] for embryonal organisms). 



Our approach was to design 

a rotation device which 

(a) can be used together with 

a wide range of commercially 

available microscopes 

[5], (b) extends the possibilities 

of various 3D microscopy 

techniques, e.g. light sheet 

fluorescence microscopy, 

confocal microscopy and 

structured illumination microscopy 

and (c) is suitable 

for specimens with a size of 

a few micrometers up to several 

millimeters. The whole 

rotation device is mounted 

on a holder that is inserted 

into the positioning stage of 

a microscope. It is helpful for 

recording images or stacks 

of images from any desired 

direction – whether for subsequent 

multi-view reconstruction 

or for just having 

a better chance to pick out 

the most interesting part of 

the sample for observation or 

imaging [6]. 

Sample Holding 

The crucial part for sample 

rotation is the way of sample 

holding. The sample has 

to be freely rotatable providing 

optimum optical access 

from every direction. In the 

presented setup (fig. 1) samples 

are located in round capillaries 

coupled to a stepping 

motor. The round capillary is 

placed in another rectangular 

capillary which is fixed. The 

outer rectangular capillary 

is made of borosilicate glass 

(n = 1.47) and its plane surfaces 

assure optimum illumination 

and image quality. 

Importance of Index 

Matching 

To prevent optical distortion 

in illumination and detection, 

the refractive index 

of the inner round capillary 

has to match the refractive 

index of the immersion fluid 

which fills the space between 

the outer and the inner capillary 

as well as the medium 

surrounding the sample. 

For fixed samples: If the 

samples are fixed in glycerol, 

round capillaries made of borosilicate 

glass are a good 

choice since the refractive indices 

are almost equal. In this 

case glycerol should be chosen 

as immersion fluid, too. 

For living samples: Samples 

located in an aqueous 

medium like agarose are 

best hold in round capillaries 

made of fluorinated ethylene 

propylene (FEP, n = 1.34) 

[7]. In this case water is used 

as an immersion fluid between 

the two capillaries. Alternatively, 

the FEP capillary 

may be used without a surrounding 

rectangular capillary 

when observing the sample 

with a water immersion 

objective lens. In that case, it 

is sufficient to couple the FEP 

capillary and the lens with a 

drop of water and to use the 

rectangular capillary only as 

a retainer at the open end of 

the FEP capillary. 

A wide range of sizes for 

rectangular and round capillaries 

are commercially 

available (e.g. VitroTubes by 

VitroCom Inc., USA). Thus, 

the user is free to choose the 

size of the capillary matching 

the size of the specimen to 

be observed. We commonly 

use outer rectangular capillaries 

with an inner cross 

section of 600 µm × 600 µm 

or 900 µm × 900 µm with a 

wall thickness of 120 µm or 

180 µm [8] in combination 

with inner round capillaries 

with an outer diameter 

of 550 µm or 870 µm with 

a wall thickness of 75 µm 

or 85 µm. When observation 

with illumination wavelengths 

in the UV range is 

desired, there is also the 

possibility to use capillaries 

made of quartz glass instead 

of borosilicate glass. Suitable 

FEP capillaries are also commercially 

available from different 

suppliers (e.g. Zeus, 

Ireland). 

Quick Sample Uptake 

Sample uptake by the capillary 

is very easy to perform 

using the capillary forces, if 

the sample is located in a liquid. 

If the sample is located 

in a gel like agarose it can be 

taken up by plunging the capillary 

directly into the agarose. 

Alternatively, in both cases the 

capillary can be attached to 

any kind of syringe or pump to 

soak in the sample. 

Observing the Sample 

The stepping motor rotating 

the inner round capillary is 

driven by a stand-alone control 

unit offering different operation 

modes including a PC 

interface. Via the control unit 

rotation speed, angular resolution 

(from 0.1125° to 1.8°) 

and direction of rotation can 

be chosen. 

We Create Solutions. 

At Applied Scientific Instrumentation 

Whether it is a complete system 

for a complex biological experiment, 

automation devices for increasing throughput, or 

inspection systems to catch defects and increase 

production, Applied Scientific Instrumentation has 

the products, people, and partners to provide 

well-engineered solutions for you. 

FOR MORE INFORMATION... 

Visit: www.asiimaging.com 

Email: info@asiimaging.com 

Call: (800) 706-2284 or (541) 461-8181 

Visit us at the 10th Forum of Neuroscience (FENS) 

Stand #128, July 2-6, 2016, Copenhagen, Denmark 



For observation a microscope objective 

lens with an appropriate working 

distance should be chosen to reach 

at least the level of the rotation axis of 

the sample. Good results can be achieved 

by objective lenses with low numerical 

apertures, e.g. 5x/0.15, 10x/0.30 or 

20x/0.50 lenses. Alternatively, for higher 

magnification a 63x/0.9 water dipping 

objective lens turned out to be suitable. 

Exemplary Results 

Figures 2 and 3 show some exemplary 

results. The images depicted are z-projections 

from eight different rotation 

angles. The copepod in figure 2 was incubated 

with rhodamine 6G at a concentration 

of 10 µM for 24 h. Fluorescence 

images were recorded by confocal 

laser scanning microscopy using an excitation 

wavelength of 488 nm. Fluorescence 

was detected using a long pass filter 

with cut-off wavelength at 505 nm. 

Image stacks from each direction consist 

of 100 images recorded at distances 

of ∆z = 6 µm. 

Figure 3 shows z-projection images of 

a zebrafish embryo recorded by confocal 

laser scanning microscopy at the Institute 

of Molecular Biology (IMB), Mainz, 

Germany. 

References 

All references are available online: 

http://bit.ly/IM-Bruns 

Affiliation 

1 

Aalen University, Institute of Applied 

Research, Aalen, Germany 

Contact 

Dr. Thomas Bruns 

Aalen University 

Institute of Applied Research 

Aalen, Germany 

thomas.bruns@hs-aalen.de 

www.hs-aalen.de 

Fig. 2: Fluorescence z-projection images of 8 

individual rotation steps of a copepod with 

half-filled egg sac incubated with rhodamine 6G 

(10 µM, 24 h) recorded by confocal laser scanning 

microscopy (excitation wavelength: 488 nm; 

fluorescence detected at λ ≥ 505 nm; each 

z-stack: Δz = 6 µm, 100 images). 

Fig. 3: Fluorescence z-projection images of 8 

individual rotation steps of a zebrafish embryo 

recorded by confocal laser scanning microscopy 

[Images recorded by Holger Dill, Mária Hanulová, 

and Sandra Ritz, Institute of Molecular 

Biology (IMB), Mainz, Germany]. 

Read more about 3D Imaging: 

http://bit.ly/IM-3D 

Recent information on 

Confocal Laser Scanning Microscopy: 

http://bit.ly/IM-CLS 

[1] 


http://bit.ly/IM-Bruns 

Visit us at Optatec, Frankfurt · Booth C29 

OPTICAL FILTERS 

For Fluorescence Spectroscopy 

AHF analysentechnik AG · +49 (0)7071 970 901-0 · info@ahf.de 

www.ahf.de 


sCANNING PROBE MICROSCOPY 

The Multimeter at the Nanoscale 

Charge Transport at the Nanoscale Measured by a Multi-Tip Scanning Probe Microscope 

Bert Voigtländer 

A multi-tip scanning tunneling microscope (STM) 

specifically designed for charge transport measurements 

at the nanoscale is described. This versatile 

tool gives insight into fundamental transport 

properties at the nanoscale. We exploit the 

capabilities of the instrument by measuring resistance 

profiles along freestanding GaAs nanowires, 

by the acquisition of nanoscale potential 

maps, and by the identification of an anisotropy 

in the surface conductivity at a silicon surface. 

Introduction 

Fig. 1: Photo of the multi-tip scanning probe microscope with an outer diameter of only 50 mm, leading 

to highest stability. 

Since microelectronics evolves into nanoelectronics, 

it is essential to perform electronic 

transport measurements at the nanoscale. 

The standard approach to this is 

to use lithographic methods for contacting 

nanostructures. However, in research 

and development stages other methods 

to contact nanoelectronic devices may be 

more suitable. An alternative approach for 



Fig. 2: (a) Schematic of a four-point measurement on a nanowire. (b) SEM 

image of a freestanding nanowire contacted by three tips. The STM tips act like 

the test leads of a multimeter, however, contacting objects at the nanoscale. 

Fig. 3: Potential map measured on a Si surface with the main potential drop 

occurring at the atomic step edges. The current flows from the top to the 

bottom in this image (image size 0.5 µm). 

Fig. 4: Anisotropy of the surface conductivity of the Si(111)-7x7 surface. (a) 

Schematic of the square four-point configuration with the steps on the 

surface indicated by the diagonal lines. (b) Four-point resistance measured 

as function of the rotation angle. The resistance is low if the current is 

directed parallel to the step edges, while it is large when the current is 

perpendicular to the step edges. 

the contacting of nanostructures is to use 

the tips of a multi-tip scanning probe microscope, 

in analogy to the test leads of a 

multimeter used at the macroscale. The 

advantages of this approach are: (a) Flexible 

positioning of contact tips and different 

contact configurations are easy to 

realize, while lithographic contacts are 

permanent. (b) in situ contacting of “as 

grown” nanostructures still under vacuum 

allows to keep delicate nanostructures 

free from contaminations which can be induced 

by lithography steps performed for 

contacting. (c) Probing with sharp tips can 

be non-invasive (high ohmic), while lithographic 

contacts are invasive (low ohmic). 

In order to use a scanning tunneling 

microscope (STM) [1] for electrical measurements 

at nanostructures, more than 

one tip is required. Thus, we developed 

an ultra-stable multi-tip instrument 

which gives access to the above outlined 

advantages in nanoprobing [2]. This is in 

accord with the recent paradigm shift in 

scanning probe microscopy which transforms 

from “just imaging” to extended 

measurements at the nanoscale. 

Multi-Tip Microscope 

The instrument (fig. 1) comprises four 

scanning units allowing for a completely 

independent motion of all four tips. A 

scanning electron microscope (SEM) image 

of the four tips brought close to the 

sample under study is shown in the eyecatcher 

image of this article. Imaging 

with the secondary electrons leads to a 

shadow effect (dark shadow image of the 

tip apex) giving access to the tip sample 

distance. Recently, a startup company 

[4] has been founded which offers this 

instrument. In the following, we demonstrate 

the capabilities of the instrument 

for nanoscale charge transport measurements 

by presenting some examples. 

Resistance Profiling along Nanowires 

As a first example, we present nanoscale 

resistance mapping along freestanding 

GaAs nanowires with a diameter of ~100 

nm [5], which are still “as grown” upright 

and attached to the substrate. The schematics 

in figure 4a, and in an SEM image 

in figure 4b shows three tips brought into 

contact with a nanowire, realizing a fourpoint 

resistance measurement. 

In the STM based approach of nanocontacting, 

four-point measurements 

can be performed not only in one single 

configuration, as it is the case for the 

lithographic approach, but many configurations 

can be measured by moving 

the tips along the nanowire (a movie of 

this can be accessed in the web [4]). In 

this way we can measure a resistance 

profile along the nanowire with 50 data 

points. 

Potential Maps 

Another method giving valuable insight 

into the charge transport properties 

of nanostructures is the scanning tunneling 

potentiometry (STP). Nanoscale 

potential maps are acquired during the 

flow of electrical current. Implementing 

STP in a multi-tip setup has several advantages 

[6]. The potential resolution is 

Read more about 

scanning probe microscopy: 

http://bit.ly/IM-SPM 

More detailed version of this article: 

http://bit.ly/multi-tip-spm 

[1] 


http://bit.ly/IM-Voigtlaender 


a couple of µV. We have applied 

the STP technique on Si 

surfaces (fig. 3) and could determine 

the surface conductivity 

on the terraces as well 

as the step resistivity [6]. 

Anisotropic Conductance 

The increasing importance 

of surface conductance 

compared to conductance 

through the bulk in modern 

nanoelectronic devices calls 

for a reliable determination 

of the surface conductivity. 

A model system for corresponding 

investigations 

is the Si(111)-7x7 surface. 

The challenge is to disentangle 

the contribution due 

to the surface conductivity 

from the bulk conductivity. 

We have developed a method 

which uses distance dependent 

four-probe measurements 

in the linear configuration 

in order to determine 

the surface conductivity [7]. 

Moreover, also the anisotropy 

of the surface conductivity 

can be measured by the 

four-probe method, when the 

tips are arranged in a square 

arrangement and are rotated 

(fig. 4(a)). In the current case 

the anisotropy is induced by 

a parallel arrangement of 

atomic steps on the surface. 

The continuous behavior of 

the measured four-point resistance 

as function of the 

rotation angle is shown in 

figure 4b. From these data 

the step resistivity as well as 

the resistivity of the terraces 

can be determined [7]. 

Success 


A multi-tip scanning tunneling 

microscope can be 

like a multimeter at the nanoscale 

in order to contact 

nanostructures by the tips 

and performing subsequently 

electrical measurements. This 

multi-tip based approach of 

nanoprobing has the advantage 

of a very flexible probe 

(re-) positioning, allowing for 

many different probing geometries 

on a single nanostructure. 

Moreover, contaminations 

of the nanostructures 

inherent to the lithographic 

approach are avoided and the 

probing contacts can be noninvasive. 

Altogether, the SPM 

based nanoprobing approach 

allows to perform a large variety 

of nondestructive electrical 

measurements at the 

nanoscale. Currently, the instrument 

is developed further 

towards a multi-tip AFM/STM 

to allow for an improved performance 

on partly insulating 

samples. 

References 


online: http://bit.ly/ 

IM-Voigtlaender 

When innovation and technology align 

Contact 

Dr. Bert Voigtländer 

Peter Grünberg Institut (PGI-3) 

Forschungszentrum Jülich 

Jülich, Germany 

b.voigtlaender@fz-juelich.de 

www.mprobes.com 

AFM 

Asylum Research 

“ 

My AFM from 

Asylum Research 

allowed me to 

move fast in the 2-D 

materials field.” 

Andras Kis 

Associate Professor 

EPFL Switzerland 

Conclusion and Outlook 

Professor Kis will be speaking at Euro AFM Forum, 

University of Geneva, 22-24 June 

For more information: www.oxford-instruments.com/EuroForum 

AFM.info.eu@oxinst.com 

+49 612 2937 0 

www.oxinst.com/AFM


Integrated Raman – FIB – SEM 

A Correlative Light and Electron Microscopy Study 

Frank Timmermans 1 ,Barbara Liszka 1 ,Derya Ataç 2 , Aufried Lenferink 1 , Henk van Wolferen 3 , Cees Otto 1 

Fig.1: Raman microscope objective integrated in the FIB-SEM vacuum chamber: (Left) Raman microscope objective integrated in the FIB-SEM (FEI NOVA 

Nanolab 600) vacuum chamber. (Right) Raman microscope added onto the FIB-SEM vacuum chamber. 

We present an integrated confocal Raman microscope 

in a FIB - SEM. The integrated system 

enables correlative chemical specific Raman, 

and high resolution electron microscopic analysis 

combined with FIB sample modification on 

the same sample location. New opportunities in 

sample analysis using correlative Raman-SEM, 

and Raman – FIB – SEM are demonstrated on 

different samples in materials and biological 

sciences. 

chamber, with the other components positioned 

onto and outside the electron 

microscope, as presented in figure 1. 

The integration places no limitation on 

the operation of either the Raman or the 

FIB-SEM. Figure 1 (left) shows both the 

Raman objective and integrated 3D XYZ 

stage, used for sample scanning during 

optical microscopy. 

Introduction 

The field of integrated Correlative Light 

and Electron Microscopy (iCLEM) has 

witnessed an enormous growth over 

the last decade. Different optical microscopes 

have been integrated in electron 

microscopes, with the integration performed 

by both commercial and scientific 

organizations [1, 2]. In this article 

a Raman microscope integrated with a 

focused ion beam (FIB) – scanning electron 

microscope (SEM) system is presented. 

The commercial optical Raman 

microscope, from HybriScan Technologies 

B.V., is specifically designed for integration 

in the SEM vacuum chamber. 

It functions as an add-on module bringing 

the optical objective into the vacuum 

Fig. 2: Correlative high resolution SEM (A) and chemical specific Raman (B) analysis of multiple crystals, 

and crystal polymorphisms. (C) Specific sample structures are identified with SEM, and the corresponding 

Raman spectra are shown in (D, E, and F). Chemical specific Raman spectroscopy is used for 

compound identification, showing: calcium sulfate (location 1, 2, 3, 4, 5), the calcium carbonate polymorphism 

vaterite (location 6), the calcium carbonate polymorphism calcite (7), and multiple fluorescence 

spectra from the photosynthetic bacteria M. aeruginosa [3]. 


PIONEERS BY 

PROFESSION 

The Mars rover Spirit began automated analysis 

of the planet‘s rocks and soil in January 2004. 

• Absolute excitation Laser power 

determination in 0.1 mW steps (TruePower). 

• High-res spectroscopy and 

imaging in parallel 

• User-friendly access to industryleading 

performance 

3D confocal Raman image of 

CCl4 in an oil-water-alkane emulsion 

apyron 

Automated Raman Imaging 

The WITec apyron Raman system automates the exploration 

of sample surfaces and chemical components. Software 

controlled, self-adjusting measurement routines along with 

TruePower and motorization bring a revelation in userfriendliness 

to cutting edge Raman imaging. 

Feel the spirit of exploration, automatically. 

WITec apyron – convenience meets performance. 

Raman • AFM • SNOM • RISE 

apyron – 

Winner of the 

ACHEMA Innovation 

Award 2015 

WITec Headquarters 

WITec GmbH, Lise-Meitner-Str. 6, 89081 Ulm, Germany, phone +49 (0)731 140700, info@witec.de 

www.witec.de


Fig. 3: (A) SEM image of multiple graphene flakes, 

noticeable by the darker color for multilayers. (B) 

Raman spectrum measured on the position indicated 

in A, of a representative location on the 

graphene flake, the D, G, and 2D bands are indicated. 

(C, D, E) integrated Raman intensity of the D, 

G, and 2D graphene bands. 

Chemical Specificity 

with Electron Microscopy 

Integrated Raman and electron microscopy 

enables the correlative analysis of 

samples with both chemical specific Raman- 

and nanometer resolution electron 

microscopy. Applications demonstrating 

correlative chemical specificity and high 

resolution are performed on multiple 

samples. First a sample containing multiple 

different crystals is analyzed, showing 

spectral Raman analysis correlated 

to particle morphologies observed with 

SEM. Second a sample of graphene flakes, 

fabricated through chemical deposition, 

is investigated for potential contaminations 

and spectral intensity analysis of 

the D, G, and 2D Raman bands. Correlative 

chemical and high resolution microscopic 

analysis is demonstrated in figure 

2, where crystal sub-micron morphological 

and chemical specific analysis is demonstrated. 

The samples contain many different 

structures two calcium carbonate 

polymorphisms, calcite and vaterite and 

calcium sulfate crystals, further the photosynthetic 

bacteria M. aeruginosa is analyzed. 

Figure 2A, and B shows the correlative 

SEM and Raman cluster image 

from the observed region of interest. Specific 

structures in the analyzed region 

are indicated in 2C, and the corresponding 

spectra are presented in figure 2D, E, 

and F. Calcium sulfate crystals are identified 

by their 1006 cm -1 Raman band, and 

a needle like shape is observed in the SEM 

analysis, the polymorphisms calcite and 

Fig. 4: (A) SEM image of FIB patterned silicon. (B) 1 st order silicon Raman band analysis. (C, D, E) Intensity, 

spectral width, and peak position of the 1 st order silicon band over the FIB patterned region. (F) Raman 

band of amorphous silicon, indicated on the 450 cm -1 region. (G) Intensity map of amorphous silicon [3]. 

vaterite are identified by the Raman band 

positions at 1086 cm -1 and 1088 cm -1 

respectively. 

Correlative Raman Electron 

Microscopy of Graphene 

Correlative Raman micro-spectroscopy 

with electron microscopy is performed on 

a sample containing graphene flakes (fig. 

3). The sample is fabricated with a chemical 

vapor deposition method on a nickel 

substrate. The process fabricates singleand 

multi-layer graphene, with multi-layers 

visible as darker areas in SEM analysis. 

The graphene structure quality on 

nickel is investigated with Raman microspectroscopy. 

The known Raman bands 

for graphene the 2D, G and D bands are 

visible in the spectra. The graphene D 

band is often an indication of disorder in 

the graphene structure. Performing a Raman 

microscopic image reveals an overall 

high quality sample, low D-band intensity, 

specific the locations with high 

D-band intensity can potentially be further 

investigated at higher resolution using 

correlative electron microscopy. Further 

the Raman intensity maps of the 

G- and 2D- band are provided in figure 3D 

and E, showing increased Raman activity 

for multi-layer graphene on nickel. The 

use of Raman spectroscopy for detection 

of single or multiple layers of graphene 

and analysis of the thickness uniformity 


http://bit.ly/IM-Raman 

More information on Focused Ion 

Beam: http://bit.ly/IM-FIB 

[1] 


http://bit.ly/IM-Timmermans 



and spatial distribution of the 

flakes is demonstrated, additional 

correlative SEM enables 

the high resolution analysis of 

interesting sample features or 

potential defects. Furthermore 

electron microscopy enables 

the fast localization of regions 

of interest (ROI) for chemical 

specific Raman analysis. Further 

applications, for example, 

using correlative Raman 

analysis after FIB modification 

of graphene are within reach 

using the correlative FIB-SEM- 

Raman microscope [3]. 

tion has placed on limitation 

on operation of the FIB, SEM 

or Raman microscope, thus 

FIB modification and SEM 

or Raman microscopic analysis 

of large samples e.g. 6 

inch wavers is possible. Correlative 

Raman microscopy 

in-situ in the vacuum chamber 

is enabled and demonstrated 

on multiple samples 

in combination with both FIB 

and SEM. The combination 

of Raman chemical specificity 

with FIB and SEM, as part 

of the broader field of iCLEM, 

promises exciting new opportunities 

in both biological and 

materials sciences. 

References 


online: http://bit.ly/ 

IM-Timmermans 

Affiliations 

1 

Medical Cell Biophysics 

group, MIRA institute, University 

of Twente, Enschede, 

The Netherlands 

2 

NanoElectronics group, 

MESA+ institute, University 

of Twente, Enschede, The 

Netherlands 

3 

Transducers Science and 

Technology, MESA+ institute, 

University of Twente, Enschede, 

The Netherlands 

Contact 

Frank Timmermans 

Medical Cell Biophysics group 

MIRA institute 

University of Twente 

Enschede, The Netherlands 

f.j.timmermans@utwente.nl 

www.utwente.nl/tnw/mcbp/ 

Correlative Raman 

Analysis with FIB Ablation 

Raman spectroscopy is a 

promising tool for correlative 

analysis in combination with 

FIB sample modification. Using 

the FIB for material ablation 

enables micromachining 

of samples, by ablation of 

sample surface material with 

a high energy ion beam. This 

method potentially leaves the 

sample vulnerable for contamination 

through redeposition 

of removed material, and for 

sample damage, and molecular 

defects through ion penetration 

into the sample. Raman 

analysis of a FIB patterned 

sample is demonstrated on a 

silicon waver sample (fig. 4). 

The FIB is used to pattern an 

easily recognizable structure, 

which is subsequently analyzed 

with Raman microscopy. 

The analysis reveals changes 

in the 1 st order silicon crystal 

Raman band on regions where 

FIB patterning is performed. 

Rigorous analysis enables the 

detection of peak shifts and 

band broadening with 0.1 cm -1 

accuracy. Further a broad Raman 

band at 450 cm -1 is indicative 

for amorphous and 

micro-crystalline silicon [4] 

which is accurately detected 

after FIB treatment. 

Conclusions 

The compact commercial 

Raman microscope is integrated 

as an add-on module 

to the FEI Nova Nanolab 

600 FIB-SEM. The integra-


Spectra of Electrons Emerging from PMMA 

Monte Carlo Simulation of Electron Energy Distributions 

Maurizio Dapor 

This work describes a Monte Carlo 

algorithm which appropriately 

takes into account the stochastic 

behavior of electron transport in 

solids and treats event-by-event all 

the elastic and inelastic interactions 

between the incident electrons and 

the particles of the solid target. The 

energy distributions of secondary 

and backscattered electrons emerging 

from polymethylmethacrylate 

(PMMA) irradiated by an electron 

beam are simulated and compared 

to the available experimental data. 

The Spectrum 

When an electron beam impinges 

on a solid target, many 

electrons can be backscattered, 

after they interacted 

with the atoms and electrons 

of the target. A fraction of 

them conserves their original 

kinetic energy, having suffered 

only elastic scattering 

collisions with the atoms of the 

target. These electrons constitute 

the so-called elastic peak, 

or zero-loss peak, whose maximum 

is located at the energy 

of the primary beam. Close to 

the elastic peak, another feature 

can be observed: it is a 

broad peak collecting all the 

electrons of the primary beam 

which suffered inelastic interactions 

with the outer-shell 

atomic electrons (plasmons 

losses, and inter-band and intra-band 

transitions). Another 

important feature of the electron 

energy spectrum is represented 

by the secondaryelectron 

emission distribution, 

i.e., the energy distribution of 

those electrons that, once extracted 

from the atoms by inelastic 

collisions and having 

travelled in the solid, reach 

the surface with the energy 

sufficient to emerge. The energy 

distribution of the secondary 

electrons is mainly 

confined in the low energy region 

of the spectrum, typically 

well below 50eV [1]. 

The Monte Carlo Algorithm 

The results presented in this 

paper were obtained using 

differential and total elastic 

scattering cross sections 

calculated utilizing Mott theory 

[2], i.e. numerically solving 

the Dirac equation in a 

central field; this procedure 

is known as the “relativistic 

partial wave expansion 

method” and it has been demonstrated 

to provide excellent 

results when compared to experimental 

data. On the side 

of the energy losses, the inelastic 

mean free paths are 

calculated by taking into account 

the inelastic interactions 

of the incident electrons with 

atomic electrons, phonons, 

and polarons. The calculation 

of the electron-electron inelastic 

scattering processes was 

performed within the Mermin 

theory [3]. Electron–phonon 

interactions were described 

using the Fröhlich theory [4]. 

Polaronic effect was modeled 

according to the law proposed 

by Ganachaud and Mokrani 

[5]. Electron trajectories follow 

a stochastic process, with 

scattering events separated 

by straight paths having a distribution 

of lengths that follows 

a Poisson-type law. Once 

the step length is generated, 

the elastic or inelastic nature 

Fig. 1: Energy distribution of the electrons emerging from PMMA with 

energies between 0 and 20eV. Monte Carlo simulated spectrum (red solid 

line) is compared to the Joy et al. experimental spectrum [8] (black line). 

Data are normalized to a common maximum. The primary energy is 1000eV. 

The primary electron beam is normal to the surface. Electrons are accepted 

over an angular range from 36° to 48° integrated around the full 360° 

azimuth. The zero of the energy scale is located at the vacuum level. 



© molekuul.be / Fotolia.com 

Fig. 2: Monte Carlo simulated spectrum of electrons emerging from PMMA 

with energies between 750eV and E0=800eV (E0 = primary electron energy). 

The plasmon-loss peak is located at about 22eV from the elastic peaks. The 

primary electron beam is normal to the surface. Electrons are accepted over 

an angular range from 0° to 90° integrated around the full 360° azimuth. The 

zero of the energy scale is located at the vacuum level. 

Fig 3: Monte Carlo simulated total electron yield σ = δ + η of PMMA as a 

function of the primary electron kinetic energy E0 (solid line). The Monte 

Carlo data were obtained integrating the curves of energy distribution 

including all the electrons emerging with energy from 0 to E0. Experimental 

data: taken from reference [9] (red circles) and from reference [10] (green 

triangles). 

of the next scattering event, 

the polar and azimuthal angles, 

and the energy losses, 

are all sampled using the relevant 

cumulative probabilities 

according to the usual Monte 

Carlo recipes [6]. Details of the 

Monte Carlo calculations can 

be found in reference [7]. 

Results 

The Monte Carlo energy distribution 

of the electrons 

emerging from PMMA irradiated 

by an electron beam with 

primary energy E 0 =1000eV 

is presented in figure 1. The 

spectrum is simulated assuming 

that the primary electron 

beam is normal to the surface. 

The zero of the energy 

is located at the vacuum level. 

In the same figure, a comparison 

of the Monte Carlo simulated 

spectrum to the Joy et 

al. experimental electron energy 

distribution [8] is shown. 

The same conditions of the 

experiment are used for the 

simulation, i.e. acceptance 

angles in the range from 36° 

to 48° integrated around the 

full 360° azimuth (Cylindrical 

Mirror Analyzer geometry). 

The Monte Carlo calculation 

describes very well the initial 

increase of the spectrum, the 

energy position of the maximum, 

and the full width at 

half maximum. Monte Carlo 

simulation is not able, on the 

other hand, to describe the 

fine structure of the peak, 

in particular the observed 

shoulder on the left of the 

maximum. 

In figure 2 the spectrum of 

the electrons emerging close 

to the elastic peak (backscattered 

electrons) is presented. 

The plasmon loss peak is located 

at about 22eV from the 

zero loss peak. 

In the experiments, on the 

one hand, the secondary electron 

emission yield δ is measured 

as the integral of the 

energy distribution of all the 

emitted electrons over the energy 

range from 0 to 50eV. 

The backscattering coefficient 

η, on the other hand, 

is measured integrating the 

distribution of all the emitted 

electrons over the energy 

range from 50eV to the energy 

of the elastic peak. 

In figure 3 the Monte 

Carlo simulated total electron 

emission yield σ = δ + η 

is compared, in the primary 

electron energy from 0 to 

1500eV, to the available experimental 

data [9,10]. 

Conclusion 

We have described and used 

a Monte Carlo algorithm for 

the evaluation of the energy 

distributions of secondary 

and backscattered electrons 

from polymethylmethacrylate 

irradiated by an electron 

beam. The integration of the 

distributions over the appropriate 

energy ranges allows 

calculating the secondary 

electron yield, δ, the backscattering 

coefficient, η, and 

the total electron yield, σ, 

as a function of the primary 

electron energy. The Monte 

Carlo simulated data are in 

agreement to the available 

experimental data. 

Acknowledgments 

Warm thanks are due to 

Diego Bisero (University of 

Ferrara), Giovanni Garberoglio 

(ECT*-FBK, Trento) and 

Cornelia Rodenburg (University 

of Sheffield) for fruitful 

discussions and stimulating 

suggestions. This work was 

supported by Istituto Nazionale 

di Fisica Nucleare (INFN) 

through the Supercalcolo 

agreement with FBK. 

Contact 

Dr. Maurizio Dapor 

European Centre for Theoretical Studies 

in Nuclear Physics and Related 

Areas (ECT*-FBK) 

Trento Institute for Fundamental 

Physics and Applications (TIFPA-INFN) 

Povo, Trento, Italy 

dapor@ectstar.eu 

www.ectstar.eu 

www.tifpa.infn.it 

More information on Monte 

Carlo methods in microscopy: 

http://bit.ly/IM-MC 

Read more about the microscopy 

of PMMA: http://bit.ly/IM-PMMA 

References: 

[1] http://bit.ly/IM-Dapor 



Stemming Unwanted Interference 

Resolution Improvement by Incoherent Imaging with ISTEM 

Florian Krause 

In Transmission Electron Microscopy (TEM) spatially incoherent image formation can have significant 

advantages regarding attainable resolution by removing unwanted interference effects. This 

has been exploited in the scanning TEM mode, which is incoherent but limited by other factors. 

Combining a scanning beam with the conventional TEM imaging mode can overcome these limitations. 

This method called ISTEM gives access to the advantages of both modes and facilitates an increase 

in resolution. 

From Traditional TEM to ISTEM 

High resolution Transmission Electron 

Microscopy (TEM) is one of the most important 

tools for the investigations of nanoscale 

structures. Historically, it has 

mostly been divided into two modes: 

For Conventional TEM (CTEM) the 

specimen is illuminated with a plane 

electron wave and then the image is 

formed by the objective lens of the microscope. 

For modern field emission sources 

the image formation is almost completely 

coherent here. Because a large area is illuminated, 

CTEM is influenced neither by 

the positioning precision of the incoming 

beam nor by aberrations of the probe 

forming lenses. Another advantage is the 

fact that though the size of the electron 

source has an influence on the images, it 

is not the factor limiting the resolution. 

Due to the coherence however, highresolution 

CTEM images can show complex 

interference patterns and hence be 

difficult to interpret. The high coherence 

also causes a strong dependence of the 

image pattern on the energy of the incident 

electrons. Chromatic aberration is 

therefore the limiting factor for resolution 

in CTEM. 

In the Scanning TEM (STEM) mode, 

the electron beam is focused onto the 

specimen. Then the intensity in a specific 

area of the diffraction pattern is 

recorded with an extended, usually circular 

or annular, detector. The image is 

formed by scanning over an area of the 

specimen. It can be shown that STEM is 

effectively an incoherent imaging mode 

due to the universal principle of reciprocity 

[1]. Therefore it is much more ro- 


Someone has to be first. 

XFlash ® FlatQUAD, multiple detector systems & VZ 

Analyze textured 

samples with ease. 

The unique XFlash ® FlatQUAD offers a large solid angle and an amazing take-off angle. 

This minimizes shadowing so you can see every nook and cranny of your sample in record time. 

Bruker‘s multiple detector systems provide large solid angles and great take-off angles too. 

You can start with one detector building up to four, depending on your needs. 

The Variable Z (VZ) adapter allows you to optimize take-off angles in-situ, which significantly 

improves the analysis of topographically challenging samples. 

Someone has to be first. 

www.bruker.com/quantax-flatquad 

Innovation with Integrity 

EDS


Fig. 1: Principle of ISTEM: (A) For each scan point of the STEM illumination the objective lens creates an 

image from the electrons leaving the specimen in a small area (circle). (B) If the camera acquires over 

the entire scanning process, all the images from the scan positions are added up incoherently and very 

little interference can occur. The actual path of the electron beam is not of importance for this (arrows). 

bust towards chromatic aberration [2], 

while interpretation of its image patterns 

is much more straightforward compared 

to CTEM. However, it is intuitive to see 

that the resolution in STEM is limited 

by the size of the focused probe, which 

is proportional to the size of the electron 

source. A second limitation to the STEM 

imaging is the precision with which the 

electron probe can be positioned during 

the scan process. 

The central idea of ISTEM, which 

stands for Imaging STEM, is to combine 

CTEM and STEM to get the best of both 

modes [3]: For this the focused scanning 

STEM beam is used to illuminate the 

specimen, while like in CTEM the objective 

lens is used to create an image. 

probe at each time only a very small spot 

of the specimen is illuminated. The objective 

lens then creates an image in the 

camera plane. When at a later time another 

point is illuminated, again an image 

is formed. Because of the different times 

no interference between both scan points 

can occur. If the camera is set to acquire 

for the entire time the STEM beam needs 

to scan over the area of interest, all scan 

points add up incoherently and there is 

very little interference. Only specimen 

points that are simultaneously illuminated 

by the probe can interfere. From 

this intuitive description it becomes clear 

that spatially incoherent image formation 

can be realized with ISTEM. 

In detailed wave optical calculations it 

can even be shown that the images taken 

with ISTEM do not depend on the aberrations 

of the probe forming lenses at all 

[4]. Costly aberration correction of the 

probe is therefore not necessary for the 

used microscopes. 

Similarly the electron source size is 

not of importance. Furthermore, looking 

at figure 1 it can also be understood that 

the precision of the beam positioning is 

also not relevant: As long as the scan area 

is homogeneously filled with STEM beam 

positions it does not matter whether the 

path of the electron probe is actually a 

straight line or a zigzag course, since unlike 

in STEM the image is directly formed 

by the objective lens and an unprecise 

beam position does only shift the illumination 

but not the image. ISTEM thus has 

all the advantages of CTEM. 

Due to the incoherence of ISTEM, it 

can also be expected to share the related 

benefits with STEM and indeed figure 2 

clearly demonstrates that the effect of increasingly 

strong chromatic aberration 

is much smaller in ISTEM images, which 

do not change much at all, than for 

CTEM, where resolution is quickly lost. 

In similar studies it can also be demonstrated 

that, like STEM, ISTEM high resolution 

images are in general relatively 

simple compared to the more complex 

CTEM patterns. Therefore ISTEM can legitimately 

be said to combine all the advantages 

of CTEM and STEM. 

Overcoming Conventional Limits 

Realization of Incoherence 

The schematic principle of ISTEM is illustrated 

in figure 1: Thanks to the focused 

Advantages of ISTEM 

Because chromatic aberration is the primarily 

limiting factor in modern microscopes 

for CTEM, the robustness towards 

it, which was demonstrated in figure 2, 

shows ISTEM‘s potential to overcome it. 

Fig. 2: CTEM (left) and ISTEM images (right) of diamond in [110] projection for increasing chromatic 

aberration characterized by the defocus spread. The structure is still resolved for large spreads in 

ISTEM. 

Fig. 3: ISTEM image of GaN in [1-100] projection. 

The N and Ga atomic columns are distinctly 

resolved as also can be seen in the line scan. 

Read more about 

Scanning TEM: 

http://bit.ly/S-TEM 

Basic information on TEM: 

http://bit.ly/TEM-basiv 

[1] 


http://bit.ly/IM-Krause 



The maximum resolution that 

can be reached with a TEM 

is referred to as the information 

limit. It is usually measured 

with a Young fringe experiment, 

which even tends to 

overestimate the point resolution 

that is actually possible. 

The FEI Titan employed 

for the ISTEM experiment 

presented in the following has 

an information limit of 81 pm 

at an acceleration voltage of 

300 kV and is equipped with 

an aberration corrector for 

the objective lens. It was used 

to take ISTEM micrographs of 

an 8 nm thin crystalline gallium 

nitride lamella, where 

the electron beam fell along 

[1-100] direction. In this projection, 

there are two atomic 

columns, one consisting of 

gallium and one of nitrogen 

atoms, that have a distance of 

only 63 pm. Hence they cannot 

be resolved separately 

under conventional operation 

of the used microscope. With 

ISTEM however, as figure 3 

clearly shows, the images of 

both columns are distinctly 

separated. Just changing to 

the STEM illumination hence 

indeed allows for a substantial 

improvement of resolution 

that overcomes the conventional 

information limit 

of the microscope thanks to 

its realization of incoherence. 

This is even more remarkable 

as nitrogen is much 

lighter than gallium; a fact 

that makes their simultaneous 

imaging difficult for many 

other techniques like e.g. annular 

dark-field STEM. 

It should be emphasized 

here, that, opposed to other 

realizations of incoherent illumination, 

ISTEM can in fact 

be used on every microscope 

that allows both CTEM and 

STEM operation, which is the 

case for almost all contemporary 

instruments. It does not 

require any hardware modifications 

and is not much more 

difficult in its application than 

usual CTEM operation. 

ing method for many microscopic 

applications. It can be 

shown that by an appropriate 

choice of the apertures 

ISTEM is able to yield the 

same images as most established 

STEM techniques but 

without the influence of electron 

source size or unprecise 

scanning, which again means 

an improvement of resolution. 

First experimental studies 

in this direction have 

shown encouraging results. 

Another recently proposed 

idea is the use of IS- 

TEM for the acquisition of 

energy filtered images where 

simulations prove its capability 

to suppress unwanted 

artefacts [5]. In conclusion, 

the ISTEM method allows 

pushing the point resolution 

of electron microscopes well 

beyond their usual limits by 

a combination of the two traditional 

modes realizing incoherent 

imaging. 

‘ 

Acknowledgment 

The author thanks the EMAT 

in Antwerp and the ERC in 

Jülich for fruitful cooperation. 

References 

All references available online: 

http://bit.ly/IM-Krause 

Contact 

MSc Florian Krause 

University Bremen 

Institute of Solid State PhysicsBremen, 

Germany 

f.krause@ifp-uni-bremen.de 

Future Prospects 

With the presented advantages 

ISTEM is a promis-


Electro-Optical Characterization of 3D-LEDs 

Nondestructive Inspection of 4’’ Wafers in Bird’s Eye View by an FE-SEM 

Johannes Ledig, Sönke Fündling, Frederik Steib, Jana Hartmann, Hergo-Heinrich Wehmann, Andreas Waag 

The optimization of three dimensional LEDs with core-shell geometry requires adapted characterization 

methods with high spatial resolution. Integrating manipulators with small probe tips inside 

a cathodoluminescence scanning electron microscope (CL-SEM) enables the investigation of local 

electro-optical properties without the need for elaborate contact preparation. Moreover this allows 

for precise monitoring of the contact position by SEM imaging and to correlate electroluminescence 

and CL measurements. 

Introduction 

Three dimensional light emitting diodes 

(3D-LEDs) with a core-shell geometry 

are supposed to have substantial advantages 

over conventional planar LEDs 

[1–3]. The active area along the sidewalls 

of the structures can considerably 

be increased by high aspect ratios - 

leading to a lower current density inside 

the InGaN multi quantum well (MQW) 

at the same operation current per substrate 

area. Such LEDs were recently developed 

within the frame of the EU-FP7 

funded project GECCO and the DFG research 

group FOR1616. The production 

of devices out of arrays of these 3D-LEDs 

grown by metalorganic vapor phase epitaxy 

(MOVPE) is already scaling up to 

substrates with larger areas [4], generating 

a request for reliable characterization 

techniques of local electro-optical 

properties with high spatial resolution 

on different positions along the substrate. 

As also subsequent device processing 

should be performed on a wafer 

scale the applied techniques need to be 

non-destructive. 

For this purpose, an electron microscope 

equipped with a field emission 

gun (FEG) and a large specimen 

chamber capable for full stage scanning 

of 4-inch wafers also in bird’seye 

view has been installed in 2015 at 

the epitaxy competence center (ec²) of 

Braunschweig University of Technology. 

Optical characterization inside the 

view field of the SEM is possible by a 

CL setup, which consists of a parabolic 

mirror inserted below the pole piece 

and a spectrograph attached to the system. 

The chamber is actively isolated 

from vibrations and piezo controlled 

manipulators are mounted on the sample 

stage. This combination enables 

precise mechanical manipulation monitored 

by SEM imaging as well as electrical 

and electro-optical characterization 

of nanostructures. Using a triax cabling 

for the electrical probes, also high impedances 

(e.g. single nanostructures) 

can be analyzed in a two- or three-point 

configuration. 

Configuration Details 

The system is based on a Tescan Mira3 

GMH FE-SEM including scintillator 

based detectors for secondary electrons 



(ET-type SE and In-Beam SE) and backscattered 

electrons (motorized low-kV 

BSE) (fig. 1). The electron beam absorbed 

current (EBAC) as well as the 

electron beam induced current (EBIC) 

between two contacts can be measured 

and imaged by an integrated detector. 

Precise electrical contacting is performed 

using Kleindiek MM3A-EM manipulators 

equipped with low current 

measurement kits (LCMK). Beside these 

detectors for electron related signals a 

Gatan MonoCL4 CL-setup is attached 

to the microscope chamber to monitor 

photon-related response from the sample. 

Its parabolic collection mirror was 

designed on request for investigation of 

planar samples at tilt angles up to 30 °. 

Due to the FEG a small electron 

probe spot can be achieved in the optical 

focus point at a working distance 

of 10 mm, even with beam energies of 

only a few keV. This enables a high spatial 

resolution also in BSE, CL and EBIC 

imaging, although probing of 3D-structures 

is influenced by scattering and 

shadowing of signals in the ensemble. 

At such conditions, electrical and optical 

properties of the sample, e.g. the 

gradient (fig. 2) or fluctuations in the 

pn-junction and InGaN QW along the 

sidewall [2,5], can be probed with a 

high spatial resolution. 

The opening figure presents a color 

overlay of the SE (red) and EBIC (cyan) 

image of an ensemble of InGaN / GaN 

core-shell LEDs visualizing the light 

emitting region of the center structure 

contacted by a tungsten probe tip. 

Optical Detection 

The optical spectrometer setup is 

equipped with different diffraction gratings 

and a CCD camera for parallel detection 

of a whole spectrum in a single 

shot as well as a photomultiplier (PMT) 

for fast band pass detection of luminescence, 

both covering a broad spectral 

range from the UV to the NIR. The 

PMT is used for fast mapping of CL excitation 

images, in particular for subsequent 

capturing stacks of monochromatic 

images taken at different 

wavelength. Such stacks are used for 

evaluation of optical properties with a 

high spatial resolution; arbitrary band 

pass images can be generated also by 

post processing or by optical filters (fig. 

2). The SEM is able to grab up to four 

signals simultaneously during the full 

pixel dwell time (starting at 20 ns). A 

drift correction of slices in the stack can 

also be applied afterwards by correlating 

corresponding SE images. However, 

with respect to the small drift of samples 

fixed by clamping this is usually 

not necessary. CL spectra probed by exciting 

a small region of the sidewall reveal 

a gradient of the InGaN/GaN MQW 

emission along the height (fig. 2). This 

CL also includes defect related yellow 

luminescence (YL) and near band edge 

© Sergey Nivens | Fotolia 

STAY UPDATED 

G.I.T. LAB PORTALS 

Receive all relevant information on 

the laboratory business and academic 

research in chemistry and biotechnology 

with G.I.T. Newsletters. 

Subscribe for free at: 

laboratory-journal.com/user/register 

GIT-labor.de/user/register (German Edition) 

Imaging-GIT.com/user/register 

Fig. 1: Colorized photo of the FE-SEM setup highlighting the 4” LED wafer (cyan) at a working distance 

of 10 mm tilted by 30 °, manipulator (green), special parabolic mirror for light collection (blue, partly 

retracted), low-kV BSE (magenta, partly retracted), pole piece with In-Beam SE (orange) and SE detector 

(red). 



Fig. 3: Stereographic SE image generated by tilting the beam by ±0.2° on a 

24 µm field of view of an ensemble of InGaN/GaN core-shell LEDs obtained 

with 10keV and at a sample tilt of 30°; one structure is contacted by a 

tungsten probe tip. The anaglyph can be viewed by red-cyan glasses. 

Fig. 2: (a) Spectra of CL and EL obtained by exciting small areas at the top 

part of the sidewall by a 590 pA electron beam of 10 keV and by driving a 

current of 1.5 µA from the probe tip contact to a buffer contact, respectively. 

(b) CL mapping of the InGaN MQW emission using a 500 nm short 

pass filter. 

emission (NBE) from the GaN 

outside the active region. 

Tilting the Electron Beam 

The electron optics (EO) of 

this FE-SEM is also capable of 

rocking the electron beam in a 

cone of up to ±12° for mapping 

of electron channeling patterns 

(ECP) on a small area of 

about 15 µm in diameter. Such 

ECP are also used to align the 

sample (via stage tilt and rotation) 

in specific diffraction 

conditions of the crystal struc- 

ture to evaluate the density of 

threading dislocations and its 

type by electron channeling 

contrast imaging (ECCI) [6]. By 

switching from the BSE detector 

to the mirror for light collection 

also a correlative analysis 

of ECCI with EBIC and CL 

can be performed at the same 

diffraction condition. 

This EO tilt can also be 

used to image the sample by 

the SEM from a certain direction 

without affecting the tip 

contact by stage movement. 

A subsequent scanning of the 

sample from different incident 

directions enables topography 

reconstruction and generates 

a three dimensional impression, 

e.g. by a stereographic 

image as given in figure 3. 

Beam Blanking for 

Electrical Measurements 

Beside in situ measurements 

the point contacts are used to 

obtain the IV-characteristics 

and also electroluminescence 

(EL) utilizing the CL-setup 

while blanking the electron 

beam (fig. 2). Care is given to 

arrange the probe tip for contacting 

in the position of the 

optical focus, neither touching 

nor significantly shadowing 

the mirror and sample. 

Due to lack of a contact layer 

the current is crowding locally 

at the contact [5]. 

EL spectra evolve already 

at currents of a few nA locally 

driven through the point contacts 

on small InGaN/GaN LED 

structures. The MQW emission 

of EL is shifted compared 

to CL which is assigned to 

the different types of excitation 

and might also be related 

to an inhomogeneous MQW 

stack. No significant current 

spreading occurs along the 

p-type GaN shell as it has a 

lower conductivity than the 

n-type region; hence the EL 

is mainly originating from a 

small volume close to the contact. 

Subsequent contacting at 

different positions therefore 

gives a sub-µm spatial resolution 

of local electro-optical 

properties and the gradient of 

the InGaN composition along 

the sidewall facet can also be 

revealed by EL spectra [1,7,8]. 

Acknowledgements 

The financial support of the 

MWK, DFG and BMBF is 

highly acknowledged. 

References 


online: http://bit.ly/IM-Ledig 

Affiliation 

Braunschweig University of 

Technology, Institute of Semiconductor 

Technology and 

Laboratory for Emerging Nanometrology, 

Braunschweig, 

Germany 

Contact 

Dipl.-Ing. Johannes Ledig 

Braunschweig University of Technology 

Institute of Semiconductor Technology, 

epitaxy competence center ec² and 

Laboratory for Emerging Nanometrology 

Braunschweig, Germany 

j.ledig@tu-braunschweig.de 

www.tu-braunschweig.de/iht 

Read more about microscopy 

of light emitting diodes: 

http://bit.ly/IM-LED 

Further information on 

cathodoluminescence microscopy: 

http://bit.ly/CL-SEM 

[1] 


http://bit.ly/IM-Ledig 


STAY UPDATED WITH 

G.I.T. NEWSLETTERS 

LABORATORY-JOURNAL.COM 

GIT-LABOR.DE 

© fotogestoeber | Fotolia 

IMAGING-GIT.COM 

Subscribe for free at: 

www.laboratory-journal.com/user/register 

www.GIT-labor.de/user/register (German Edition) 

www.Imaging-GIT.com/user/register 

Receive all relevant information on the laboratory business and academic 

research in chemistry and biotechnology with G.I.T. Newsletters.

Products 

USB 3.0 Range Expanded 

Jenoptik’s Progres Gryphax 

Arktur and NAOS cameras 

are set to be the latest additions 

to the Company’s USB 

3.0 product range. A further 

three cameras for scientific 

applications will complete the series at the end of the year. The 

models will complement the range of cameras for routine use 

in microscopy. With a high light yield and wide dynamic range, 

the micro universe can be explored at a very low noise level on 

high-resolution monitors. Thanks to the USB 3.0 interface live 

images are presented at video speed, and users can experience 

the microscope’s performance capabilities on the monitor without 

delay. Positioning samples and focusing is now faster. The 

wide range of applications includes medical usage, quality and 

material controls, and forensics. Images can be captured with a 

resolution of between two and eight megapixels (MPix). 

Near Field Scanning Optical Microscope 

Mad City Labs has introduced the MCL-NSOM, 

a versatile near field scanning optical microscope 

(NSOM). This aperture NSOM is 

built on the manufacturer’s RM21 inverted 

optical microscope, which allows 

users to convert to aperture-less 

NSOM, AFM and fluorescence optical 

microscopy. It includes XYZ 

closed loop nanopositioning for 

sample and fiber scanning providing 

sub-nanometer precision and low noise performance. 

Six axes of automated positioning are provided, including independent 

automation for fiber alignment to the optical axis, 

and are controlled with Labview based software. Included are 

a fiber launch with excitation source, alignment camera, and 

detection avalanche photodiode (APD) but it can also easily accommodate 

user ancillary excitation and detection optics. 

Jenoptikwww.jenoptik.com 

Mad City Labs 

www.madcitylabs.com 

Nine Structures in One Imaging Measurement 

LED Light Source for Fluorescence Imaging 

The X-Cite 120LEDBoost by Excelitas 

is a broad spectrum LED 

light source for fluorescence imaging 

applications. It is replacing 

the X-Cite 120LED: offering 

the same key features and mercury-free 

benefits as its predecessor, 

it delivers 40% more power for improved fluorophore 

excitation. Designed with intuitive controls and no bulbs or 

modules to install, set-up and operation is simpler. The light 

source provides high-power, broad-spectrum fluorescence excitation 

from 370-700nm, catering to a wide range of popular 

fluorophores and fluorescent proteins. The 40% power increase 

in the 500-650nm region of the spectrum provides stronger excitation 

for fluorescent proteins such as mCherry and other red 

fluorophores. Ergonomic fingertip control, very fast PC control 

and TTL triggering provide multiple control options to limit 

photobleaching and maximize flexibility. 

In collaboration with Picoquant, Profs. 

Markus Sauer (Julius Maximilian University 

of Würzburg, Germany) and 

Jörg Enderlein (Georg-August-Universität, 

Göttingen, Germany) have 

published a novel strategy for Fluorescence 

Lifetime Imaging (FLIM) for 

visualizing nine different target molecules 

at the same time. This method 

was realized with the Microtime 200 microscopy platform and 

the Symphotime 64 data acquisition and analysis software. 

FLIM makes it possible to visualize structures and processes in 

cells by exciting molecules with light and employing the lifetime 

information of the triggered fluorescence. The newly devised 

approach, named spectrally resolved FLIM (sFLIM), uses three 

lasers with different wavelengths pulsing in an alternating pattern 

for exciting the molecular labels. As these labels exhibit 

subtle differences in their emission spectra and fluorescence 

decay patterns, the collected data can be analyzed through a 

software algorithm to distinguish them precisely. 

Excelitaswww.excelitas.com 

Pico Quant 

www.picoquant.com 

Live Cell Imaging 

Phase Focus as launched “Livecyte”, a complete live cell imaging 

system for kinetic cytometry. It is designed to enable cell 

biology researchers to achieve a deeper understanding of the 

phenotypic and kinetic behaviors of both individual and populations 

of cells in a label free environment and, will allow 

users to interleave fluorescence measurements during different 

drug treatments or across cell types within a single experiment. 

With the ability to image multiple regions of interest 

within wells and across well plates, this intuitive system delivers 

multi parametric, quantitative data and high contrast, 

high fidelity images and videos. The ability of the system to image 

continuously ensures that large fields of view can be captured 

without stitching: highly motile cells are not lost during 

long time-courses. Cell perturbation is minimized since cells 

are imaged in their growth media without the need for fluorescent 

labels. 

Phasefocuswww.phasefocus.com 


Products 

Compact Linear Positioning Stage 

The compact linear stage L-509 

from PI (Physik Instrumente) 

is providing high precision linear 

motion with minimized 

runout. It is designed for research 

and industrial applications, 

including beamline systems, microscopy, semiconductor 

manufacturing, and photonics instrumentation. Despite its compact 

dimensions, the stage can handle payloads of more than 

20 lbs. Available with travel ranges of 26mm (1”), 52mm (2”), 

and 102mm (4”) and velocity to 20mm/sec, the stage delivers 

high accuracy and smooth motion with unidirectional repeatability 

down to 0.1 microns. Long service life and very high guiding 

accuracy with minimum backlash are ensured by the precision 

crossed roller bearings with anti-creep cage assist. For 

very high precision requirements, a linear encoder with 1 nanometer 

resolution is available. 

Confocal Laser Scanning 

Microscope 

Olympus’ Fluoview FV3000 confocal 

laser scanning microscope 

combines high-performance imaging 

capabilities with ease of use so researchers in such fields 

as cell biology, cancer research and stem cell research can collect 

relevant imaging data quickly and easily. Built for fast, stable 

and accurate measurements of biological reactions within 

living cells and tissues, the microscope offers flexibility for all 

live-cell imaging applications, providing high-resolution images 

of structures and dynamic intracellular processes. It is controlled 

by an intuitive software interface so even novice users 

can generate high-quality data and images. The instrument’s 

optical design offers macro to micro imaging capabilities with 

objectives ranging from 1.25X to 150X magnification. One version 

comes with galvanometer scanner, and the FV3000RS is 

equipped with a hybrid resonant / galvanometer scanner. 

Physik Instrumente 

www.physikinstrumente.com 

Olympuswww.olympus-lifescience.com 

Dichroics for Super-Resolution Microscopy 

AHF analysentechnik offers 

now the new Semrock 

beamsplitter series 

additionally to their existing 

superflat dichroic 

program for super-resolution 

microscopy: λ /10 P-V 

per inch flatness on 3 mm 

thick dichroics and improved 

λ /2 P-V per inch 

flatness on improved 1 

mm dichroics are now available. There will be no compromise 

regarding guaranteed steepest edges, short wavelength 

reflectivity down to 350 nm, and long wavelength transmission 

optimized out to 1200 nm or 1600 nm. Super-resolution 

imaging systems are highly sensitive to optical wavefront distortion 

and demand the highest quality components. Laser 

dichroic beamsplitters with λ /10 flatness minimize the reflected 

wavefront distortion, thereby maximizing both the 

signal and the signal-to-noise ratio in super-resolution microscopes. 

1 mm thick laser dichroic beamsplitters have been 

significantly improved to λ /2 flatness (~255 m radius of curvature). 

They will fit into microscopy filter cubes and improve 

the performance of laser based confocal and TIRF illumination 

systems. They are also ideal for reflection of imaging 

beams in conventional structured-illumination techniques as 

well as patterned illumination systems for localized photoactivation. 

These dichroic beamsplitters allow the use of 

much larger diameter illumination beams, offering researchers 

and instrument developers more flexibility in system design 

with no compromise to overall performance. Please ask 

AHF for a demo system. 

WEBINAR 

FLUORESCENCE LIFETIME IMAGING 

USING FLUORESCENT DECAY RATES 

TO IDENTIFY INDIVIDUAL FLUOROPHORES 

JUNE 28, 2016 AT 2PM 

TO REGISTER FOR THIS FREE 

WEBINAR PLEASE GO TO 

bit.ly/Webinar-PCO 

AHF analysentechnik 

www.ahf.de 


Products 

Imaging Live Cells under Near-Native Conditions 

Leica Microsystems launches the Leica DFC9000, a monochrome 

microscope camera with a highly sensitive third-generation 

sCMOS sensor. The camera enables researchers to image 

live cells under near-native conditions, allowing them to 

gain a better understanding of cellular processes and dynamics. 

The sensor with its high quantum efficiency over the entire 

spectrum of light, provides a high signal-to-noise ratio to securely 

detect even faint signals. Compared to the second generation 

sensor, the maximum quantum efficiency increased by 

14%, totaling up to 82% depending on wavelength. In combination 

with a very low noise level, this results in a crisp fluorescence 

signal against a dark background. The high sensitivity of 

the camera eliminates the need to monitor GFP-overexpressing 

specimens and protects cells from phototoxicity. 

Leicawww.leica-microsystems.com 

Light Sheet Microscopy 

Andor’s sCMOS camera can be 

used for a broad range of microscopy 

applications, including 

wide field fluorescence 

microscopy, calcium ratio imaging 

and Super Resolution 

Microscopy. The Company points out that their Neo 5.5 sCMOS 

camera is particularly useful regarding latest Light Sheet Microscopy 

developments: A particular micro fabrication technique used 

to produce a mirror array that both directs the excitation beam 

and holds the sample may revolutionize the field of 3D super-resolution 

microscopy, according to an international group of scientists. 

They demonstrated that their Single-Objective Selective- 

Plane Illumination Microscopy (soSPIM) allows researchers to 

examine the activity of single proteins or entire embryos on their 

existing microscope systems. This cannot be handled by standard 

microscope systems. 

Andorwww.andor.com 

EM-Tec Silicon Nitride TEM Support Membranes 

Micro to Nano reveals its next generation 

EM-Tec silicon nitride support films for 

TEM with interesting and innovative improvements. 

To greatly improve handling, 

the silicon support frame has been 

shaped into a 3.05 mm compatible hexagon 

with micro-structured TrueGrip edges. 

One side of the hexagon includes a reference 

notch to for sample location during processing and loading. 

Chemistry has been further improved to produce stress-optimised 

films, resulting in robust and ultra-planar silicon nitride 

films. Manufacturing of the EM-Tec silicon nitride films include 

a proprietary cleaning process to deliver clean, debris free films. 

Available with several windows sizes and membrane thicknesses 

of 20, 50 and 200nm. 

Correlative Raman-SEM Imaging 

The WITec RISE microscopy mode for correlative 

Raman-SEM imaging is now compatible 

with the scanning electron microscope Zeiss 

Merlin. The integration of both techniques 

into one system greatly improves ease-of-use 

and accelerates the experimental workflow. It 

places both the objective and sample stage required 

for Raman microscopy within the SEM’s vacuum 

chamber. Thus the sample can remain under vacuum for 

both measurements and is simply transferred between the Raman 

and SEM measuring positions by a software-driven pushbutton 

mechanism using an extremely precise scan stage. The 

combined system provides all functions and features of a standalone 

Zeiss SEM and a WITec confocal Raman microscope. 

Micro to Nano 

www.microtonano.com 

WITecwww.witec.de 

Win the book! 

To have a chance of winning the book find the original figure in this issue from 

which the image below is taken. Send the title of the article to contact@imaginggit.com 

with the subject line Read & Win! All correct answers will be entered in 

a prize draw and the lucky winner will receive a copy of “Handbook of 

Fluorescence Spectroscopy and Imaging”, which is featured on page 13. 

Closing date: 17. August 2016 

High-Speed Camera Series 

Each model of the IL5 High- 

Speed 5MP Camera series 

from Fastec Imaging is easily 

mounted on microscopes, 

enabling to record high-speed 

video of microscopic events. 

Both spatial and temporal 

magnification work in tandem to clarify understanding in applications 

such as microfluidics, where particles often move through 

the field of view very quickly. Four different models revealing resolution 

and frame rate from 2560 x 2080 @ 230fps to 800 x 600 

@ 1650fps are available. Each type record over 3200 fps at VGA 

resolution and more than 18,000 fps at smaller resolutions. Able 

to save images to an SSD or SD card while recording high-speed 

bursts of hundreds or even thousands of images at a time, the 

camera is always ready for the next high-speed snapshot. The 

IL5 can be controlled over Gigabit Ethernet via Fastec FasMotion 

software on PC/Mac or via the built-in web interface with a web 

browser on PC, Mac, tablet, and smartphone. 

Fastec Imaging 

www.fastecimaging.com 


indEX / IMPRINT 

AHF Analysentechnik 30, 49 

Aalen University 28 

Andor 23, 50 

Applied Scientific Instrumentation 29 

Argolight 24 

Asylum Research 33 

Braunschweig University of Technology 44 

Bruker micro CT 

Outside Back Cover 

Bruker Nano 41 

Digital Surf 43 

Edmund Optics 25 

European Molecular Biology Laboratory 9 

Excelitas 13, 48 

Fastec Imaging 50 

Forschungszentrum Jülich 31 

Hamamatsu Photonics 18 

Jenoptik48 

Julius-Maximilians-University of Würzburg 13 

Leica 50 

Mad City Labs 48 

Märzhäuser 19 

MCO Congres Marseille 10 

Micro to Nano 50 

Microscience Microscopy Congress 14 

Molecular Devices 

16, Cover 

NKT Photonics 

Inside Front Cover 

Olympus 49 

PCO AG 11, 12, 49 

Phasefocus48 

Physik Instrumente 7, 49 

Pico Quant 5, 48 

Piezosystem9 

Schneider Kreuznach 27 

Select Biosciences 9 

Spanish Portuguese Meeting 

for Advanced Optcial Microscopy 11 

Tescan37 

Trento Institute for Fundamental Physics 

and Applications 38 

University of Bremen 40 

University of Antwerpen 15 

University of Sheffield 21 

University of Twente 34 

Witec 35, 50 

Imprint 

Published by 

Wiley-VCH Verlag GmbH & Co. KGaA 

A Company of John Wiley & Sons, Inc 

GIT VERLAG 

Managing Directors 

Sabine Steinbach, Philip Carpenter 

Publishing Director, Chem & Lab 

Dr. Heiko Baumgartner 

Regional Commercial Director 

Dr. Katja Habermüller 

Editorial Director int. Publications 

Laboratory & Biotechnology 

Editor-in-Chief 

Dr. Martin Friedrich 

Tel.: +49 6201 606-715 · Fax: -793 

E-mail: martin.friedrich@wiley.com 

Editors 

Dr. Thomas Matzelle 

Tel.: 0151-11006677 · Fax: -793 

E-mail: thomas.matzelle@wiley.com 

Dr. Birgit Washburn 

Tel.: +49 6201 606-760 · Fax: -793 

E-mail: birgit.washburn@wiley.com 

Dr. Arne Kusserow 

Tel.: +49 6201 606-732 · Fax: -793 

E-mail: arne.kusserow@wiley.com 

Dr. Martin Graf-Utzmann 

Tel.: +49 6201 606-766 · Fax: -793 

E-mail: martin.graf-utzmann@wiley.com 

Dr. Ralf Kempf 

Tel.: +49 6201 606-751 · Fax: -793 

E-mail: ralf.kempf@wiley.com 

Editorial Assistance 

Yadigar Aksoy 

Tel.: +49 6201 606-752 · Fax: -793 

E-mail: yadigar.aksoy@wiley.com 

Isabel Brenneisen 

Tel.: +49 6201 606-716 · Fax: -793 

E-mail: isabel.brenneisen@wiley.com 

Wiley GIT Reader Service 

65341 Eltville / Germany 

Phone: +49 6123 9238 246 

Fax: +49 6123 9238 244 

E-Mail: WileyGIT@vuservice.de 

Our service is available for you from 

Monday to Friday 8 am – 5 pm CET 

Segment Sales Manager 

Laboratoty & Biotechnology 

Vanessa Winde 

Tel.: +49 6201 606-721 · Fax: -793 

E-mail: vwinde@wiley.com 

Media Consultant 

Dr. Stefanie Krauth 

Tel.: +49 6201 606-728 · Fax: -793 

E-mail: stefanie.krauth@wiley.com 

Production Manager 

Christiane Potthast 

Tel.: +49 6201 606-741 · Fax: -793 

E-mail: christiane.potthast@wiley.com 

Sales Administrator 

Claudia Vogel 

Tel.: +49 6201 606-758 · Fax: -790 

E-mail: claudia.vogel@wiley.com 

Reprints 

Dr. Stefanie Krauth 

Tel.: +49 6201 606-728 · Fax: -793 

E-mail: stefanie.krauth@wiley.com 

Advisory Board 

Prof. A. Diaspro, Italian Institute of Technology 

Dr. C. Durkan, Univ. of Cambridge, UK 

Dr. M. Dürrenberger, Univ. of Basel, Switzerland 

Dr. R. Fleck, NIBSC, Herts, UK 

Prof. M. Gu, Swinburne Univ., Australia 

Prof. B. Hecht, Univ. of Wuerzburg, Germany 

Prof. M. Hegner, Trinity College Dublin, Ireland 

Prof. F.-J. Kao, Nat. Sun Yat-Sen Univ., Taiwan 

Prof. N. Kruse, Univ. of Brussels, Belgium 

Prof. D. Nicastro, Brandeis Univ., MA, USA 

Dr. J. Rietdorf, MicroImaging Labs, Munich, Germany 

Dr. P. Schwarb, FMI, Basel, Switzerland 

Dr. D. Spitzer, ISL, France 

Prof. G. A. Stanciu, Univ. of Bucharest, Romania 

Prof. G. Valdré, Univ. of Bologna, Italy 

Dr. Roger Wepf, ETH Zurich, Switzerland 

Dr. T. Zimmermann, ORG, Barcelona, Spain 

Printed by 

pva, Druck und Medien, 76829 Landau 

Circulation 

18,000 copies, 4 times per annum 

Advertising price list from October 1 st 2014 

Subscription 2016 

Four issues 38.00 €, single copy 14.70 € ca. plus postage. 

Pupils and students receive a discount of 50 % at sight of a 

valid certificate. Subscription orders can be revoked within 

1 week in writing. Dispatch complaints are possible only within 

4 weeks after publishing date. Subscription cancellations are 

accepted 6 weeks before end of year. 

Bank account 

Commerzbank AG, Mannheim, Germany 

Account No. 07 511 188 00 

Routing No. 670 800 50 

BIC: DRESDEFF670 

IBAN: DE94 6708 0050 0751 1188 00 

Specially identified contributions are the responsibility of the 

author. Manuscripts should be addressed to the editorial office. 

We assume no liability for unsolicited, submitted manuscripts. 

Reproduction, including excerpts, is permitted only with the 

permission of the editorial office and with citation of the source. 

The publishing house is granted the exclusive right, with regard 

to space, time and content to use the works/ editorial contributions 

in unchanged or edited form for any and all purposes any 

number of times itself, or to transfer the rights for the use of 

other organizations in which it holds partnership interests, as 

well as to third parties. This right of use relates to print as well 

as electronic media, including the Internet, as well as databases/ 

data carriers of any kind. 

All names, designations or signs in this issue, whether referred 

to and/or shown, could be trade names of the respective owner. 

Printed in Germany 

ISSN 1439-4243 

www.imaging-git.com

SIM0216

Create successful ePaper yourself

Delete template?

Save as template?