reSolution_LNT_No1_en - Leica Microsystems

No. 01, Autumn 2012 

CUSTOMER MAGAZINE 

FOR NANOTECHNOLOGY 

reSOLUTION

EDITORIAL 

CONTENT 

Title 

Panchromatic cathodoluminescence image of 

cassiterite mineral (SnO 2 ) 

Image Credits 

Scott Wight1 , Ed Vicenzi2 , Doug Meier1 , and 

Kurt Benkstein1 1 National Institute of Standards and 

Technology 

2 Smithsonian Institution 

Source: Smithsonial National Mineral 

Collection, Preparation: Cut and polished 

with the Leica EM TXP Data Collection: 

FEI Company QuantaTM 200F SEM with 

Gatan, MonoCL4 Elite System 

2 reSOLUTION 

Dear Readers, 

It gives me great pleasure to welcome you to the Leica NanoTechnology (LNT) edition of reSolution magazine. As 

it is our first totally exclusive LNT magazine, we decided to provide information on application techniques for both 

biology and materials sample preparation. All of the articles were prepared by our customers from around the 

world and I would like to express our thanks to everyone who provided the high quality articles for this edition. 

Sharing such experiences helps to disseminate techniques and applications around our EM community. 

To provide further support for issues regarding sample preparation techniques, later in the year we will launch 

a Leica EM Sample Preparation Science Laboratory online service where you will be able to find even more 

information about applications and products specific to your needs. 

Late last year we launched three new products to enhance sample preparation in your laboratory; a new ion 

beam slope cutter, the TIC 3X, for materials SEM preparation; a critical point dryer, CPD300 - a prerequisite for 

good SEM preparation for biological and some materials samples; and an entry level high pressure freezer for 

freezing samples in tubes, the SPF. You can find more information about these instruments on our website at 

http://www.leica-microsystems.com/products/electron-microscope-sample-preparation/ 

This year we have also launched some exciting new products. A new family of coaters, the ACE range, was 

presented at the EMC meeting in Manchester UK. This new generation of coating systems continues in line 

with our development philosophy, to automate tedious processes and push forward the boundaries of sample 

preparation in line with the needs of the scientific community. 

I hope you enjoy this first LNT reSolution magazine and I look forward to your feedback. 

Happy Reading! ding! 

Best Wishes, es, 

Ian Lamswood wood 

Marketing Manager 

BIOLOGY 

Capturing neuro transmitter receptors 04 

and ion channels: 

High-resolution techniques to localize 

membrane proteins 

Biological Electron Microscopy at 07 

Durham University 

Dr. Martin W. Goldberg, Durham University, UK 

Substitutes for Uranyl Acetate in TEM 09 

Thin Section Post-Staining 

Perusing alternatives for staining applications 

for TEM thin sections 

Dry ultrathin sectioning combined with 13 

high pressure freezing/freeze-substitution 

improves retention and visualization of 

calcium and phosphorus ions prior to nucleation 

of mineral crystals within osteoblastic 

cultures 

INDUSTRY 

A Word on Cathodoluminescence 17 

Atomic Force Microscopy Study of a 19 

Stretched Impact Copolymer 

University of Wollongong Electron 21 

Microscopy Centre 

REGISTRATION 23 

IMPRINT 23

Capturing Neurotransmitter Receptors and Ion Channels 

High-resolution Techniques 

to Localize Membrane Proteins 

Daniel Althof1,2 , Akos Kulik1,3 1Department of Anatomy and Cell Biology, Institute of Neuroanatomy 

2Spemann Graduate School of Biology and Medicine, University of Freiburg, Germany 

3Department of Physiology II, University of Freiburg, Germany 

Neurotransmitter receptors and ion channels in the central 

nervous system are localized to synaptic and extrasynaptic 

membrane compartments of pre- and postsynaptic elements 

of neurons. The impact of the activation of these 

proteins on synaptic integration and regulation of transmitter 

release depends on their precise location relative to 

synapses, as well as on the density and coupling of molecules 

in microcompartments of the cells. High-resolution 

qualitative and quantitative visualization of membranebound 

receptors and ion channels is, therefore, essential 

for understanding their roles in cell communication. 

The ability of the nervous system to learn and respond to 

the environment refl ects an underlying capability of neurons 

to dynamically alter the number, type, and strengths 

of their connections. These connections, called synapses, 

are highly organized sites of contact between postsynaptic 

neurons and presynaptic terminals. The specialized synaptic 

membrane, contains a large variety of molecules such 

as receptors, ion channels, and associated structural proteins, 

whose precise subcellular organization facilitates its 

proper function. A large number of studies have provided 

evidence that the location of these proteins and their position 

relative to synapses substantially affects their functional 

roles. Neurotransmitter receptors, localized to the 

synaptic membrane of postsynaptic compartments of neurons, 

are directly exposed to released neurotransmitters. 

Consequently, they are activated in a transient manner, 

generating fast postsynaptic responses that are precisely 

time-locked to the presynaptic action potentials. In contrast, 

receptors localized to the extrasynaptic plasma membrane, 

remote from synaptic sites, are activated by spilledover 

neurotransmitters producing a tonic conductance that 

is not precisely time-locked to single presynaptic action 

potentials, but rather refl ects the whole network activity 

on a slower time scale 1 . Receptors can also be located 

presynaptically either on the extrasynaptic membrane of 

axon terminals or over the presynaptic grid where they are 

activated by neurotransmitters released by the same or by 

neighbouring boutons. Like neurotransmitter receptors, 

ion channels are also localized to the somato-dendritic 

membranes and axon terminals of neurons. Postsynaptic 

channels are generally playing a role in the integration and 

plasticity of synaptic inputs, as well as in the control of 

neuronal excitation by mediating slow inhibitory synaptic 

responses and contributing to the resting membrane potential. 

Presynaptic channels that are concentrated either at the 

presynaptic active zone or localized to the extrasynaptic 

membrane of boutons are involved in the regulation of neurotransmitter 

release, thereby playing a role in the presynaptic 

modulation of neuronal activity. It is, therefore, easy 

to understand that the same receptor and ion channel could 

fulfi ll very different functional requirements when targeted 

to different subcellular compartments of cells 1 . 

Furthermore, the impact of the activation of membrane 

proteins on synaptic integration and 

regulation of transmitter release 

critically depends on the density and 

functional coupling of receptors and 

ion channels in compartments of the 

target neurons, as well as on their location 

relative to excitatory and inhibitory 

synaptic sites. Thus, the question 

arises of how the precise subcellular 

location of these molecules can be 

determined at high resolution. 

For this purpose the following advanced 

high-resolution immunocytochemical 

methods have been widely 

used: (i) preembedding immunogold, 

(ii) postembedding immunogold, and 

(iii) sodium dodecyl sulfate (SDS)-digested 

freeze-fracture replica labeling 

(SDS-FRL) techniques. (i) In case of the 

preembedding immunogold method, 

an 0.8 nm or a 1.4 nm gold particle is 

coupled to the secondary antibodies 

Fig. 1 

BIOLOGY 

Akos Kulik Daniel Althof 

NANOTECHNOLOGY 3

BIOLOGY 

Fig. 2 

Fig. 1 & 2: Distribution and colocalization of 

GABA (B1) and Kir3.2 in dendrites of hippocampal 

cells as revealed by the SDS-digested freezefracture 

replica labeling technique. A, Immunoparticles 

for the GABA (B1) subunit were found 

in clusters (arrows) over the surface of dendritic 

shaft (Den) and spine (s) of a putative pyramidal 

cell. B, C, Double and triple immunogold labeling 

for Kir3.2 (5 nm particles; double arrows), GABA 

(B1) (10 nm; arrows), and PSD-95 (15 nm in C) 

revealed that the two proteins co-clustered in 

dendritic spines of pyramidal cells (B and C) and 

associated to the site of glutamatergic synapses 

indicated by immunoreactivity for PSD-95 (C). 

Scale bars, 200 nm 

4 reSOLUTION 

in order to facilitate proper penetration. 

Silver intensifi cation of the gold 

particles is subsequently carried out 

to produce a detectable particle size. 

This method produces non-diffusible 

labels, thus the precise site of the reaction 

and the location of the protein 

at extra- and perisynaptic sites can be 

determined. Synaptic proteins, however, 

cannot be detected using this 

method, most likely due to the inaccessibility 

of the epitopes in the synaptic 

specializations of fi xed tissues 2 . 

(ii) The postembedding immunogold 

method overcomes the problems of 

pre-embedding technique by reacting 

immunochemicals with the antigens 

exposed on the surface of the ultrathin 

sections and then detecting synaptic 

proteins with the same sensititvity as 

that for non-synaptic molecules. This 

also improves quantitative evaluation 

of the protein densities. 

However, in resin-embedded sections 

substantial proportions of proteins are 

buried and therefore not accessible 

for antibodies, limiting the detection 

sensitivity of this technique 2 . (iii) In 

SDS-FRL, the brain tissue is frozen 

with a high-pressure freezing machine (Leica EM HPM100) 

then frozen samples are freeze-fractured in a replica machine 

(Leica EM BAF060). Proteins are allocated to either 

the protoplasmic faces (P-faces) or the exoplasmic faces 

(E-faces) of plasma membranes. Molecules are immobilized 

with a thin layer of carbon (3-5 nm) followed by a 

further coating with a 2-nm-thick platinum/carbon layer for 

shadowing the membrane faces and then this material is 

strengthened with a 15 – 20 nm thick carbon deposit 3 . The 

SDS-FRL technique has two major advantages compared to 

conventional immunogold methods. 

First, the sensitivity of the SDS-FRL is considerably higher 

than that of the pre- and postembedding techniques, because 

membrane proteins are exposed on the two-dimensional 

surface of the replica (Fig. 1), making them readily 

accessible to immunoreagents. In addition, epitopes are 

denaturated by SDS, allowing antibodies known to be 

suitable for immunoblot analysis to react similarly with 

proteins immobilized on the replica membrane. Second, 

synaptic and extrasynaptic proteins can simultaneously 

be visualized and quantifi cation of immunogold density in 

membrane segments can be achieved. 

This immunocytochemical method, similarly to others, has 

limitations. First, the identifi cation of labeled morphological 

structures is diffi cult, therefore, it is necessary to use 

marker proteins to facilitate the identifi cation of fractured 

membranes 6 . Second, the separation of membrane proteins 

to P-face or E-face is unpredictable: some proteins are preferentially 

allocated to either the P-face, such as GABA (B1), 

Kir3.2 4 or the E-face, such as AMPA receptors 5 , whereas 

others, like gluRδ2 6 are localized to both faces. Thus, for 

quantitative studies, the allocation of the molecules should 

carefully be examined 7 . 

Taken together, these three immunocytochemical techniques 

provide complementary information about the cellular 

and subcellular distribution of proteins and are widely 

used for high-resolution qualitative and quantitative analysis 

of receptor and ion channel localization and colocalization 

in post- and presynaptic compartments of neurons. 

Contact 

Daniel Althof, Akos Kulik 

Department of Anatomy and Cell Biology, 

Institute of Neuroanatomy, Spemann Graduate School of 

Biology and Medicine, University of Freiburg, Germany 

Department of Physiology II, University of Freiburg, 

Germany 

akos.kulik@physiologie.uni-freiburg.de

References 

1. Farrant, M., Nusser Z. Variations on an inhibitory theme: phasic 

and tonic activation of GABA(A) receptors. 

Nature Rev Neuroscience 6(3), 215-229. (2005). 

2. Lujan, R., Nusser, Z., Roberts, J.D.B., Shigemoto, R., Somogyi, 

P. Perisynaptic location of metabotropic glutamate receptors 

mGluR1 and mGluR5 on dendrites and dendritic spines in the 

rat hippocampus. Eur J Neuroscience 8(7), 1488-1500. (1996) 

3. Fukazawa, Y., Masugi-Tokita, M., Tarusawa, E., Hagiwara, H., 

Shigemoto, R. SDS-digested freezefracture replica labeling 

(SDS-FRL), in Handbook of Cryo-preparation Methods for 

Electron microscopy, eds. Cavalier A, Spehner D, Humbel BM, 

CRC Press, New York, 567-586. (2009) 

4. Kulik, A., Vida, I., Fukazawa, Y., Guetg, N., Kasugai, Y., Marker, 

C.L., Rigato, F., Bettler, B., Wickman, K., Frotscher, M., Shigemoto, 

R. Compartment-dependent colocalization of Kir3.2containing 

K+ channels and GABAB receptors in hippocampal 

pyramidal cells. J Neuroscience 26(16), 4289-4297. (2006) 

Leica EM HPM100 

High Pressure Freezer for Cryofi xation 

of Biological and Industrial Samples 

5. Masugi-Tokita, M., Tarusawa, E., Watanabe, M., Molnar, E., 

Fujimoto, K., Shigemoto, R. Number and density of AMPA receptors 

in individual synapses in the rat cerebellum as revealed 

by SDS-digested freeze-fracture replica labeling. J Neuroscience 

27(8), 2135-2144. (2007a) 

6. Masugi-Tokita, M., Shigemoto, R. High-resolution quantitative 

visualization of glutamate and GABA receptors at central 

synapses. Curr opinion in Neurobiology 17, 387-393. (2007b) 

7. Fujimoto, K. Freeze-fracture replica electron microscopy combined 

with SDS digestion for cytochemical labeling of integral 

membrane proteins. Journal of Cell Science 108, 3443-3449. 

(1995) 

Instruments used for this sample preparation: 

Leica EM BAF060 

Freeze Fracture System 

BIOLOGY 

NANOTECHNOLOGY 5

BIOLOGY 

Biological Electron 

Microscopy 

Dr. Martin W. Goldberg, Durham University, UK 

The Electron Microscopy (EM) laboratory in the School of Biological and Biomedical Sciences at Durham University is an 

integral part of a wider facility spanning a range of advanced imaging tools (laser scanning and spinning disc confocal microscopes, 

TIRF microscopy and live cell imaging) as well as an ultra high resolution (

Such sample preparation is also amenable to immuno-gold 

labelling, so that we can use antibodies to locate cargoes 

and transporters in transit, as well as the proteins that make 

up this gateway (the NPC) (Fig.4). 

Contact 

Dr. Martin W. Goldberg, 

Durham University, UK 

m.w.goldberg@durham.ac.uk 

Instruments related to this sample preparation: 

Leica EM TP 

Automated Routine Tissue Processor 

Leica EM ACE200 

Low Vacuum Coater 

BIOLOGY 

Fig. 3: High magnifi cation transmission electron micrograph of high pressure frozen, freeze substituted yeast cell, clearly showing 

both leafl ets of the inner and the outer nuclear membranes, with ribosomes docked on the outer membrane and NPCs at points 

where the two membranes are joined. 

Fig. 4: Transmission electron micrograph of high pressure frozen, freeze substituted yeast cell immuno-gold labelled for a NPC 

protein. 

Leica EM CPD300 

Automated Critical Point Dryer 

Leica EM ACE600 

High Vacuum Coater 

NANOTECHNOLOGY 7

BIOLOGY 

Perusing alternatives for automated staining of TEM thin sections 

Substitutes for Uranyl Acetate in 

TEM Thin Section Post-Staining 

Nicole Fellner1,3 , Marlene Brandstetter1,3 , Karin Trimmel1,2 , and Dr. Guenter P. Resch1,3 1IMP-IMBA-GMI Electron Microscopy Facility, Institute of Molecular Biotechnology, Vienna, Austria 

2University of Applied Sciences, Wiener Neustadt, Austria 

3Campus Science Support Facilities GmbH, Vienna, Austria 

Nicole Fellner (left), Marlene Brandstetter (in the middle of the front row), 

Günter Resch (in the middle of the back row), Harald Kotisch (right) 

8 reSOLUTION 

Introduction 

Contrast in transmission electron microscopy 

(TEM) is mainly produced 

by electron scattering at the specimen: 

Structures that strongly scatter 

electrons are referred to as electron 

dense and appear as dark areas in the 

bright fi eld image, while structures 

which scatter fewer electrons appear 

bright (electron transparent) (Flegler 

et al., 1993). As electron scattering 

increases with atomic number, biological 

samples show hardly any inherent 

amplitude contrast in the TEM, 

as they are largely composed of light 

elements. 

To increase their contrast, electron dense stains can be 

added to the sample, the most commonly used heavy elements 

being: gold, platinum, tungsten, lead, and uranium. 

Biological specimens can be contrasted through various 

staining techniques: Particles like protein complexes or 

viruses can be embedded in heavy metal salts (negative 

staining), or the specimen can be covered with very thin 

electron-dense metal fi lms (replicas produced by shadowing). 

Cells and tissues can be infi ltrated with stain 

before embedding (Osmium tetroxide or uranyl acetate 

en bloc staining) or the ultra-thin sections are stained 

(Dykstra, 1992). The choice of reagents for the latter approach, 

called post-staining, is discussed in this article. 

The most frequently used method for post-staining is a 

twostep procedure of staining with uranyl acetate (UA), 

followed by lead citrate. Uranyl acetate is used as an 

aqueous or alcoholic solution with a pH for the saturated 

solution in the range of 3.5 to 4.0. The addition 

of alcohol, especially methanol, increases the solubility 

(Hayat, 2000). Uranyl acetate strongly stains proteins 

as well as nucleic acids and phospholipids. When 

applied after the uranyl acetate staining, lead citrate 

(prepared according to Reynolds, 1963) will increase this 

contrast (Dykstra, 1992). Staining can be performed either 

manually or automatically, both techniques have their advantages. 

For manual staining, a grid is fl oated, sectionside 

down, on a drop of a uranyl acetate solution for 10 

minutes. After blotting off the stain, the grid is rinsed 

thoroughly with water to remove any residual unbound 

stain. This fi rst step is followed by 5 min. lead citrate 

staining, following the same procedure. The consumption 

of reagents is minimal, whereas the effort is relatively 

high. Alternatively, poststaining can be automated. This 

ensures increased reproducibility and time saving, though 

the amount of reagents used is higher. Using the automated 

contrasting device EM AC20 (Leica Microsystems, 

Vienna) allows for simultaneous staining of up to 20 grids 

per run with no effort and a guarantee for safety, both for 

the environment and the user. Although UA is an excellent 

and well characterized stain, replacements are sought for, 

for several reasons. 

When it needs to be handled as a powder, it is very toxic 

and carcinogenic if inhaled. Furthermore, also depleted 

uranyl acetate is considered a radioactive material, and 

hence subject to relevant regulations. Therefore, UA requires 

adequate storage and careful handling, which in 

turn increases cost for shipping and waste disposal. To 

minimize contact, the automated version with the AC20 

is preferred by many users, in particular as readily prepared 

solutions are available, handling of solid UA can be 

avoided. Two reagents described in literature as replacements 

for UA caught our attention: oolong tea extract 

(OTE) and Platinum Blue. Only very little data is published 

about them and the methods are not well known in the 

EM community. Therefore, we have tested both with 

manual contrasting and for the fi rst time with the Leica 

EM AC20 instrument. 

To allow a direct comparison of results from the different 

post-staining techniques, the same sample was used for 

all tests: Liver tissue freshly dissected from mice was fi xed 

with 2.5% glutaraldehyde in 100 mmol/l Soerensen phosphate 

buffer and post-fi xed with 2.0% osmium tetroxide.

Pieces of tissue were dehydrated and embedded in Agar 

100 epoxy resin and sectioned to a nominal thickness of 

70 nm. Poststaining was performed as described below. 

Besides contrasting effi ciency and comparability of the 

results with UA, a number of other important parameters 

were assessed. 

Oolong Tea Extract 

OTE was described as post-stain to electron microscopy by 

Sato et al. (2003), and used in a small number of studies 

(Sato et al., 2008; Miller and Simakova, 2010). According 

to Rumpler et al. (2001), OTE is a type of half-fermented 

tea and produced as a foodstuff. Therefore it was assumed 

to be harmless for health and environment, even though 

the supplier failed to produce a material safety data sheet. 

It can be purchased from Ted Pella (http://www.tedpella. 

com), and is delivered as a powder. According to Sato et al. 

(2003), the polyphenolic components in OTE react with peptide 

bonds. A reaction with OTE and lead citrate in return, 

leads to an enhancement of the contrast. 

After tests with different OTE concentrations, 0.2% OTE 

dissolved in boiling ddH 2 O was used for further experiments, 

as already proposed by Sato et al. (2003). Compared 

to conventional staining with uranyl acetate, the manual 

application of OTE and the subsequent staining step with 

Reynold’s lead citrate was more time consuming (Table 1). 

Optimal results on the Leica EM AC20 were obtained also 

using an extended program at room temperature (Table 2). 

At a concentration of 0.2% OTE both manually as well as 

automatically stained samples show an increase in contrast 

even though it was clearly lower than with UA (Figs. 

1 and 2). In our hands, contamination was observed more 

frequently than with UA, which mainly constituted a problem 

at lower magnifi cation. Despite extended washing 

steps, both with manual staining or the Leica EM AC20, 

this problem could not be eliminated. 

Platinum Blue 

Platinum Blue (Pt Blue) was used as an alternative to uranyl 

acetate in thin section post-staining by Inaga et al. (2007; 

2009). This compound is a product of the reaction of cisdichlordiamineplatinum 

(II) with thymidine (Inaga et al., 

2007). The reagent can be ordered directly from the Japanese 

producer Nisshin (http://nisshin-em.co.jp). Pt Blue is 

a hazardous material, which may cause eye irritation, cancer 

and effects on fertility as well as severe disorders of 

the bone marrow, kidneys and the nervous system (MSDS 

Nisshin). The commercially available 6% stock solution has 

been found to give good results at a dilution of 1:100 for the 

manual staining and 1:200 for the automated procedure. 

The incubation times for the manual staining method were 

the same as for UA staining (cf. Table 1). 

For staining using the Leica EM AC20, the step for Pt Blue 

were extended to 30 min, using the identical conditions 

as for UA. The grids were then rinsed with distilled water, 

stained with lead citrate, and washed as described in 

Table 2. Figures 1 and 2 illustrate that all organelles show 

good contrast and that the results are comparable in quality 

to pictures taken from sections stained with UA. Differences 

can be found in a more intensely stained mitochondrial 

matrix and a higher contrast of glycogen granules. The 

ribosomes appear weaker stained as compared to UA. 

Stain 1 Time Washing Stain 2 Time Washing 

UA 10 min 2 min ddH 2 O Lead citrate 5 min 2 min ddH 2 O 

OTE 25 min 6 min ddH 2 O Lead citrate 5 min 2 min ddH 2 O 

Pt Blue 10 min 2 min ddH 2 O Lead citrate 5 min 2 min ddH 2 O 

Table 1: Details of manual staining procedures 

Stain 1 Time Washing Stain 2 Time Washing 

BIOLOGY 

UA 30 min 2 min 20 sec ddH 2 O Lead citrate 7 min 2 min 20 sec ddH 2 O 

OTE 40 min 5 min ddH 2 O Lead citrate 7 min 2 min ddH 2 O 

Pt Blue 30 min 2 min 20 sec ddH 2 O Lead citrate 7 min 2 min 20 sec ddH 2 O 

Table 2: Automated staining procedures of the Leica EM AC20. The conditions for UA and lead citrate are suggested by Leica and can be found in the user's manual of the staining device. 

NANOTECHNOLOGY 9

BIOLOGY 

10 reSOLUTION 

Conclusion 

OTE and Pt Blue were assessed as substitutes for UA in 

section post-staining for electron microscopy with regard 

to contrasting effi ciency, toxicity, handling, and price. Manual 

as well as automatic staining procedures with the Leica 

EM AC20 were tested. No en bloc staining was performed. 

Furthermore, resins other than Agar 100 or different section 

thicknesses may lead to different results. 

Both the rather weak contrast obtained with OTE as well 

as the contamination observed on the specimen were not 

convincing with manual staining and the Leica EM AC20. Pt 

Blue delivered clearly better results with both approaches. 

However, slight differences in contrasting properties as 

compared to UA have to be taken into account for interpretation 

of micrographs (Yamaguchi et al., 2010). The quality 

of results that can be obtained from automatic staining is 

comparable with the manual procedure for both OTE and 

Platinum Blue. 

Regarding toxicity, both reagents tested as substitutes of 

UA have the advantage of not being radioactive. Furthermore, 

working with health-damaging inhalable powder can 

be avoided. Being a food product it can be presumed that 

OTE is non-hazardous. In contrast, Pt Blue is toxic but as it 

is delivered as a stock solution further risky handling can 

be reduced to a minimum. The utilization of an automatic 

staining device such as the Leica EM AC20 can help to further 

minimize contact with toxic reagents. 

Staining with UA and Pt Blue is comparable in time 

and labor, whereas contrasting with OTE is more time 

consuming without delivering as satisfactory results. 

As with the Leica EM AC20, it is possible to stain 


Leica EM AC20 

Automatic Constrasting Instrument for Ultrathin Sections 

Leica EM TRIM2 

Specimen Trimming Device for TEM, SEM, LM 

up to 20 grids at once and all steps are carried out 

automatically, this becomes less of a disadvantage of OTE. 

From the economic point of view, OTE was by far the cheapest 

product. The price per grid at the used concentration of 

0.2% was signifi cantly lower than for Pt Blue (at a dilution 

of 1:100) and the 2.0% UA solution. 

Summing up, electron microscopists in need of a replacement 

for UA for post-staining of sections have the choice 

between one reagent at a very cheap price and minimal 

risk with moderate results – Oolong tea extract – and 

another one, that delivers very convincing results, but at 

higher cost and not without safety risks – Platinum Blue. 

The experiments here have shown that this is applicable 

for both manual as well as automated staining with the 

Leica EM AC20. 

Acknowledgements 

The authors would like to thank Yanli Tong (Leica Microsystems, 

Shanghai) for assistance with acquiring reagents and 

Jean Trichereau (IMBA, Vienna) for providing samples. The 

work of N.F., M.B. and G.P.R. was supported by the City of 

Vienna/Zentrum fuer Innovation und Technologie through 

the Spot of Excellence grant “Center of Molecular and Cellular 

Nanostructure.” 

Contact 

Dr. Günter Resch 

Head of Electron Microscopy, Campus Science Support 

Facilities GmbH, Vienna, Austria 

guenter.resch@csf.ac.at 

Leica EM UC7 

Ultramicrotome for Perfect Sectioning at Room Temperature and Cryo


1. MSDS Nisshin: http://nisshin-em.co.jp/home/msds/index. 

html. 

2. Dykstra, M.J. 1992. Biological Electron Microscopy: Therory, 

Techniques, and Troubleshooting. Plenum Press, New 

York. 

3. Flegler, S.L., Heckman, J.W., Jr., Klomparens, K.L. 1993. Scanning 

and Transmission Electron Microscopy: An Introduction. 

W.H. Freeman and Company, New York. 

4. Hayat, M.A. 2000. Principles and Techniques of Electron Microscopy: 

Biological Applications. 4 th ed. Cambridge University 

Press, Cambridge. 

5. Inaga, S., Hirashima, S., Tanaka, K., Katsumoto, T., Kameie, 

T., Nakane, H., Naguro, T. 2009. Low vacuum scanning electron 

microscopy for paraffin sections utilizing the differential 

stainability of cells and tissues with platinum blue. Arch Histol 

Cytol. 72(2):101-106. 

6. Inaga, S., Katsumoto, T., Tanaka, K., Kameie, T., Nakane, H., 

Naguro, T. 2007. Platinum blue as an alternative to uranyl acetate 

for staining in transmission electron microscopy. Arch 

Histol Cytol. 70(1):43-49. 

7. Miller, A.A., and A.V. Simakova. 2010. Application of Method 

of OTE Staining of Ultrathin Sections Based on Example of Microsporidia 

(Protozoa: Microsporidia). Cell and Tissue Biology. 

4(1):109-115. 

8. Reynolds E.S. 1963. The Use of Lead Citrate at High pH as an 

Electron Opaque Stain in Electron Microscopy. Journal of Cell 

Biology 17:208-212. 

9. Sato, S., Adachi, A., Sasaki, Y., Ghazizadeh, M. 2008. Oolong tea 

extract as a substitute for uranyl acetate in staining of ultrathin sections. 

Journal of Microscopy. 229(Pt 1):17-20. Sato, S., Sasaki, Y., 

Adachi, A., Dai, W., Liu, X.L., Namimatsu, S. 2003. Use of oolong 

tea extract (OTE) for elastin staining and enhancement in ultrathin 

sections. Med Electron Microsc. 36(3):179-182. 

10. Rumpler, W., Seale, J., Clevidence, B., Judd, J., Wiley, E., Yamamoto, 

S., Komatsu, T., Sawaki, T., Ishikura, Y., Hosoda, K. 

2001. Oolong Tea Increases Metabolic Rate and Fat Oxidation 

in Men. J. Nutr. 131: 2848–2852. 

11. Yamaguchi, K., Suzuki, K., Tanaka, K. 2010. Examination 

of electron stains as a substitute for uranyl acetate for the 

ultrathin sections of bacterial cells. Journal of Electron 

Microscopy (Tokyo) 59(2):113-118. 

BIOLOGY 

Fig. 1: Results from the manual poststaining procedure with UA, 

OTE, and Pt Blue, followed by lead citrate, in comparison. All images 

were acquired under identical conditions and are reproduced with 

similar contrast enhancement to allow a direct comparison of the 

contrast obtained. (A, top left) Unstained mouse liver tissue. (B, top 

right) Routine EM staining shows a good and consistent contrast of 

all cell organelles. (C, bottom left) Organelles including nucleus and 

rough endoplasmic reticulum are clearly seen in OTE stained tissue. 

(D, bottom right) Pt Blue stained sections show good contrast. 

Staining is more intense in mitochondria and glycogen granules as 

compared to UA. Scale bar: 1 μm. 

Fig. 2: Comparison of mouse liver sections after automatic staining 

with UA, OTE, and Pt Blue using the Leica EM AC20. (A, top left) 

Unstained tissue. (B, top right) UA stained liver tissue shows an 

overall good contrast. (C, bottom left) Contrast is weaker in OTE 

stained liver tissue and minor contamination is visible. (D, bottom 

right) Nucleus, rough endoplasmic reticulum, mitochondria, and glycogen 

granules are clearly visible in this specimen stained with Pt 

Blue (1:200). Scale bar: 1 μm. 

Note added in proof: Readers interested in 

replacing uranyl acetate are also referred to 

Nakakoshi, M., Nishioka, H., and Katayama, 

E. 2011: New versatile staining reagents for 

biological transmission electron microscopy that 

substitute for uranyl acetate. Journal of Electron 

Microscopy 60(6): 401–407. 

NANOTECHNOLOGY 11

BIOLOGY 

Dry Ultrathin Sectioning Combined With High Pressure 

Freezing/Freeze-substitution Improves Retention and 

Visualization of Calcium and Phosphorus Ions Prior to 

Nucleation of Mineral Crystals Within Osteoblastic Cultures 

Jeff P. Gorski1 , N.T. Huffman1 , T. Hillman-Marti2 , and Daniel Studer2 1Department of Oral Biology and the UMKC Center of Excellence 

in Mineralized Tissue Research, School of Dentistry, 

Univ. Missouri-KC, Kansas City, Kansas City, MO and 

2Institute of Anatomy, University of Bern, Bern, Switzerland 

12 reSOLUTION 

We have used cultured UMR106-01 osteoblastic cells to 

investigate the process of bone mineralization. UMR106- 

01 cells as well as primary calvarial bone cells assemble 

spherical extracellular supramolecular protein-lipid complexes, 

termed biomineralization foci (BMF), in which the 

fi rst crystals of hydroxyapatite mineral are deposited (Midura 

et al., 2004; Wang et al., 2004). A major difference 

between these culture models is the speed with which 

mineralization occurs, ranging from 12-16 days after plating 

for primary osteoblastic cells to 88h for UMR106-01 

cells. 

If mineralization is blocked by omission of phosphate 

source or by addition of serine protease inhibitor AEBSF, 

BMF complexes are formed but no mineralization occurs. 

Interestingly, ultra structural studies have shown that prior 

to mineralization BMF contain numerous membrane limited 

vesicles ranging in size from 50 nm to 2 microns in diameter. 

However, the fi rst mineral crystals are not detected 

until 78 h after plating of UMR106-01 cells and are localized 

within spherical sites presumed to be vesicles. 

Specifi cally, confocal Raman spectral analyses have shown 

that mineralization within BMF is a progressive, multi-step 

process occurring simultaneously in all BMF within a culture 

fl ask (Wang et al., 2009). Importantly, several protein 

spectral changes are detectable within each BMF prior 

to the deposition of poorly crystalline hydroxyapatite and 

when mineralization was blocked, these changes did not 

Jeff P. Gorski Daniel Studer 

Thérèse Hillmann 

occur. Thus, mineralization within BMF is a temporally 

synchronized process. However, understanding the biochemical 

mechanism of mineralization requires a detailed 

appreciation of calcium and phosphorus ion handling prior 

to crystal nucleation within BMF. 

Previous work has proposed that cartilage and/or bone 

mineralization utilizes either a single vesicle population 

enriched in both calcium and phosphorus, or, two vesicle 

populations separately enriched in calcium or phosphorus 

(Fig. 1) (Anderson, 1967; Bonucci, 1967; Arsenault and Ottensmeyer, 

1984). In order to clarify this issue, the solubility 

of these ions necessitates the use of additional methods 

such as high pressure freezing and freeze substitution to 

avoid loss during specimen fi xation, embedding, and sectioning. 

Since most prior studies have not consistently 

avoided water during specimen processing, the true impact 

of pseudo non-aqueous processing on the process of 

osteoblast-mediated mineralization is diffi cult to assess. 

The goal of our study was to use the synchronized UMR106- 

01 culture model to devise and validate a pseudo nonaqueous 

processing method to image the distribution of 

calcium and phosphorus ions within BMF immediately prior 

to nucleation of the fi rst hydroxyapatite crystals therein 

(Fig. 1). We believe this method should also be applicable 

to investigations of temporal changes in calcium ion distributions 

in other cells such as muscle.

Calcium and phosphorus can be lost from samples either 

after high pressure freezing and freeze-substitution or during 

conventional fi xation. To evaluate the effectiveness 

of our pseudo non-aqueous method, we chose to stop the 

cultures at 76 h, twelve hours after adding the ß-glycerol 

phosphate mineralization stimulus, but 2 h before the appearance 

of the fi rst crystals of mineral within BMF (Midura 

et al., 2004; Wang et al., 2009). Some cultures were 

randomly chosen to be processed for conventional fi xation 

(Fig. 2) while others were processed by high pressure 

freezing (Leica EM PACT) and freeze-substitution (Fig. 3). 

Comparison of these paired cultures support several conclusions. 

Large areas contain a somewhat homogeneous 

particulate organic matrix after conventional fi xation. 

These areas seem to exclude membrane limited vesicles 

from their volume. In contrast, high pressure frozen cultures 

contain numerous almost “white” extracellular areas, 

roughly 0.5-1 μm in diameter, within BMF (arrows, Fig. 3). 

In higher power views, it is evident that despite the low 

contrast these “white” regions do possess an underlying 

detail which represents a range of irregularly shaped 

spherical bodies from about 50 to 800 nanometers in diameter 

(not shown). However, the presence of these “white” 

areas raised immediate concerns regarding the loss of inorganic 

or organic substances. In particular, when compared 

with similar BMF regions from paired, conventionally fi xed 

cultures (Fig. 2), we hypothesized that “white” spots represented 

materials which were preferentially retained by 

high pressure freezing but which were subsequently lost 

upon further processing. 

Notably, subsequent electron spectroscopic imaging of 

calcium and phosphorus was not possible in either conventionally 

fi xed nor in freeze-substituted samples which were 

sectioned on water regardless of whether specimens were 

post-stained or not (results not shown). We therefore substituted 

dry sectioning for wet sectioning of high pressure 

frozen, freeze substituted cultures. Calcium and phosphorus 

retention is improved in dry sectioned high pressure frozen, 

freeze-substituted cultures. It is clear that substitution 

of dry sectioning leads to a dramatic increase in retention 

of calcium and phosphorus within biomineralization foci 

[compare Figures 3 (wet sectioning) and 4 (dry sectioning)]. 

Focal 0.5-1 μm diameter areas which appeared as “white” 

spots in Fig. 3 after wet sectioning, now appear dark (Fig. 

4A) in the zero loss energy image. The fact that UMR106-01 

cultures mineralize in a reproducible, temporally synchronous 

manner facilitates these direct comparisons among 

different cultures (Wang et al., 2009).In addition, electron 

spectroscopic imaging demonstrates that the dark appearing 

areas are enriched in calcium and phosphorus (compare 

Figs. 4A, B, and C). Finally, as shown in the overlay image 

in Fig. 4D, the calcium (red) and phosphorus (green) signals 

largely overlap each other in the 76 h cultures as shown 

by the presence of yellow. Since other studies have shown 

that 76 h UMR106-01 cultures do not contain detectable 

mineral crystals (Huffman et al., 2007; Wang et al., 2009), 

the enriched contents of calcium and phosphorus observed 

here could represent amorphous calcium phosphate (Driessens 

et al., 1978) and/or labile organic forms of phosphorus 

such as polyphosphates (Omelon et al., 2009). 

Importantly, a functional role for the observed enriched focal 

contents of calcium and phosphorus in mineralization is 

supported by analyses of un-mineralized control cultures. 

When a similar high pressure freezing, freeze substitution, 

and dry sectioning approach was applied to un-mineralized 

UMR106-01 cultures, few dark (calcium and/or phosphorus 

enriched) vesicles or particles were detected (not shown). 

Finally, an additional advantage to use of the oscillating 

knife was that it reduced compression during sectioning 

and reduced wrinkling of resultant sections. 

BIOLOGY 

Fig. 1: The single and dual vesicle models of extracellular mineralization. Upper panel, 

single matrix vesicle model. Calcium and phosphorus ions are progressively concentrated 

within a single population of matrix vesicles through the proposed actions of 

Ca +2 -pumping ATPase, Na + -phosphate co-transporter, and phosphatases acting on 

phospholipids. Once Ca +2 and phosphate ions reach a threshold concentration (yellow), 

nucleation of initial mineral crystals occurs leading to breakage of the vesicle 

and release of crystals which can propagate additional crystals within the surrounding 

extracellular collagenous matrix. Lower panel, dual vesicle model. Calcium and 

phosphorus are progressively concentrated within two different populations of vesicles 

which biochemically display distinct functional distributions including Ca +2 pumping 

ATPase and Na + -phosphate co-transporter activities, respectively. Subsequently, 

these calcium and phosphate enriched vesicle populations fuse and nucleate mineral 

crystals leading to breakage of the vesicle and release of crystals which can propagate 

additional crystals within the surrounding extracellular collagenous matrix. 

Instrument related to this sample 

preparation: 

Leica EM PACT2 

High Pressure Freezer with Rapid Transfer System 

NANOTECHNOLOGY 13

BIOLOGY 

Fig. 2: Appearance of BMF in mineralizing osteoblastic culture after conventional chemical fixation and wet sectioning. UMR106-01 cells were cultured for 12 h in the presence of ß-glycerol phosphate. Arrowhead 

defines the outlines of an extracellular BMF enriched in a somewhat homogeneous particulate organic matrix. 

Fig. 3: High-pressure frozen, freeze-substituted osteoblastic cultures contain BMF with translucent spots (arrow) which are poorly contrasted extracellular 

matrix regions. When compared with analogous samples subjected to dry sectioning, the translucent spots appear black (see Figure 4). 

14 reSOLUTION

In summary, our results show that high pressure freezing 

and pseudo non-aqueous processing are required to detect 

extracellular sites of early calcium and phosphorus enrichment 

in mineralizing osteoblastic cultures. Use of dry 

sectioning proved to be a critical step in the preservation 

of calcium and phosphorus. Also, electron spectroscopic 

imaging demonstrated that darkly stained vesicles within 

extracellular biomineralization foci are enriched in calcium 

and phosphorus prior to the detection of crystalline mineral 

(Wang et al., 2009). We now plan to use this method to 

determine if osteoblastic cells in vitro and in vivo utilize a 

single or dual vesicle mineralization mechanism (Gorski et 

al., 2004; Midura et al., 2009). 

Space does not permit us to cite all the relevant publications. 

Please refer to our recent publication: Studer et al., 

2011 for a more complete citation list. 

Address correspondence to: Jeff P. Gorski, Ph.D., Dept. of 

Oral Biology, School of Dentistry, University of Missouri-Kansas 

City, 650 East 25th Street, Kansas City, MO 64108. 

Phone: 815-235-2537; fax: 816-235-5524; gorskij@umkc.edu 

Contact 

Jeff P. Gorski, Ph.D. 

Professor 

UMKC Center of Excellence in Mineralized Tissues 

Department of Oral Biology 

School of Dentistry 

University of Missouri-Kansas City 

gorskij@umkc.edu 

Dr. sc. nat. Daniel Studer 

Institute of Anatomy, University of Bern, Bern, Switzerland 

daniel.studer@ana.unibe.ch 


1. Anderson, H.C. (1967) Electron microscopic studies of induced 

cartilage development and calcifi cation. J Cell Biol 35: 

81-101. 

2. Arsenault, A.L., F.P. Ottensmeyer (1984) Visualization of early 

intramembranous ossifi cation by electron microscope and 

spectroscopic imaging. J Cell Biol 98: 911-921. 

3. Bonucci, E. (1967) Fine structure of early cartilage calcifi cation. 

J Ultrastruct Res 20, 33-50. 

4. Driessens, F.C., J.W. van Dijk, J.M. Borggreven (1978) Biological 

calcium phosphates and their role in the physiology 

of bone and dental tissues. 1. Composition and solubility of 

calcium phosphates. Calcif Tissue Res 26: 127–137. 

5. Gorski, J.P., A. Wang, D. Lovitch, D. Law, K. Powell, R.J. Midura 

(2004) Extracellular bone acidic glycoprotein-75 defi nes 

condensed mesenchyme regions to be mineralized and localizes 

with bone sialoprotein during intramembranous bone 

formation. J Biol Chem 279: 25455–25463. 

6. Huffman, N.T., J.A. Keightley, C. Chaoying, R.J. Midura, 

D. Lovitch, P.A. Veno, S.L. Dallas, J.P. Gorski (2007) 

Association of specifi c proteolytic processing of bone 

sialoprotein and bone acidic glycoprotein-75 with mineralization 

within biomineralization foci. J Biol Chem 282: 

26002–26013. 

BIOLOGY 

Fig. 4: After pseudo non-aqueous processing, mineralized BMF were enriched in calcium and phosphorus. Cells were 

grown identically to those in Figures 2 and 3. A: Zero loss energy image depicting dark appearing vesicle. B: Electron 

spectroscopic imaging of calcium signal. C: Electron spectroscopic imaging of phosphorus. D: Superimposed images of the 

views in A-C. 

7. Midura, R.J., A. Wang, D. Lovitch, D. Law, K. Powell, J.P. 

Gorski (2004) Bone acidic glycoprotein-75 delineates the 

extracellular sites of future bone sialoprotein accumulation 

and apatite nucleation in osteoblastic cultures. J Biol Chem 

279: 25464–25473. 

8. Midura, R.J., A. Vasanii, X. Su, S.B. Midura, J.P. Gorski 

(2009) Isolation of calcospherulites from the mineralization 

front of bone. Cells Tissues Organs. 189:75-79. 

9. Omelon, S., J. Georgiou, Z.J. Henneman, L.M. Wise, B. Sukhu, 

T. Hunt, C. Wynnyckyj, D.Holmyard, R. Bielecki, M.D. 

Grynpas (2009) Control of vertebrate skeletal mineralization 

by polyphosphates. PLoS One 4: e5634. 

10. Studer, D., T. Hillman-Marti, N.T. Huffman, J.P. Gorski (2011) 

Eliminating Exposure to Aqueous Solvents Is Necessary for 

the Early Detection and Ultrastructural Elemental Analysis 

of Sites of Calcium and Phosphorus Enrichment in Mineralizing 

UMR106-01 Osteoblastic Cultures. Cells Tissues Organs 

May 30. [Epub ahead of print]. 

11. Wang, C., Y. Wang, N.T. Huffman, C. Cui, X. Yao, S. Midura, 

R.J. Midura, J.P. Gorski (2009) Confocal laser Raman microspectroscopy 

of biomineralization foci in UMR 106 osteoblastic 

cultures reveals temporally synchronized protein changes 

preceding and accompanying mineral crystal deposition. J 

Biol Chem 284: 7100–7113. 

NANOTECHNOLOGY 15

INDUSTRY 

A Word on Cathodoluminescence 

Cathy Johnson, Nanotechnology Division, Leider Lane, Buffalo Grove, IL 

Cathodoluminescence microanalysis is an emerging technique that is fast gaining popularity in the world of materials 

science. CL is a light emission phenomena resulting from the electron beam excitation of a luminescent material. As electronic 

transitions occur between the conduction and valence bands, CL photons are generated and detected. Electronic 

transitions due to defect levels within the band gap, particularly in the case of semiconductors and devices, can also 

infl uence CL data. Data acquisition results in a mapping of the optical activity for a specimen. 

Cathy Johnson, Leica Microsystems 

CL data can indicate defects such as imperfections or impurities 

within the microstructure of a material phase. These 

defects can have an effect on the material’s optical, electrical 

and mechanical properties. Utilizing the high resolution 

capability of a SEM or STEM, a spectrum can be acquired 

at each point location (i.e. hyperspectral imaging). As such, 

it serves as an important spectroscopy and imaging technique 

in the characterization of materials. Image resolution 

is dependent on instrument confi guration, experimental parameters 

and specimen interaction, but can range from < 10 

nanometers to the micron level. 

The fi rst MAS Cathodoluminescence Topical Conference 

was hosted October 24-28, 2011 by the National Institute 

of Standards and Technology (NIST) in Gaithersburg, MD. 

This conference was sponsored by the Microbeam Analysis 

Society (MAS), and was co-sponsored by the Australian 

Microbeam Analysis Society (AMAS). The four day program 

included a pre-conference tutorial targeted for the CL 

novice on October 24 th . The remaining three days included 

a combination of technical presentations, hands-on laboratory 

demonstrations and a contributed poster session. Presentation 

topics included: CL theory, data quantifi cation, 

advances in instrumentation, analysis and databases. Applications 

in geological, semiconductor and nanomaterial 

disciplines including sample preparation and Correlative 

CL in conjunction with complementary techniques such as 

EBIC and EBSD were also addressed. 

Instruments rel related to this sample preparation: 

Leica EM TXP 

Target Surfacing System 

Leica EM TIC 3X 

Ion Beam Slope Cutter 

Leica EM RES101 

Ion Milling System 

Contact 

Cathy Johnson 

Leica Microsystems 

Nanotechnology Division 

1700 Leider Lane 

Buffalo Grove, IL 60089 

Cathy.Johnson@leica-microsystems.com 

Panchromatic cathodoluminescence image of cassiterite mineral (SnO 2 ) 

Image Credits 

Scott Wight 1 , Ed Vicenzi 2 , Doug Meier 1 , and Kurt Benkstein 1 

1 National Institute of Standards and Technology 

2 Smithsonian Institution 

Source: Smithsonial National Mineral Collection, Preparation: Cut 

and polished with the Leica EM TXP Data Collection: FEI Company 

QuantaTM 200F SEM with Gatan, MonoCL4 Elite System

Atomic Force Microscopy 

Study of a Stretched Impact 

Copolymer 

INDUSTRY 

Dalia G. Yablon, Jean Grabowski, and Andy H. Tsou, Exxon Mobil Research and Engineering, Clinton, New Jersey, USA. 

Abstract 

Atomic force microscopy (AFM) is a powerful tool in 

the suite of nanoscale characterization techniques that 

provides a variety of information including topography, 

mechanical properties, and electrical properties with nanoscale 

lateral and sub-nanometer vertical resolution. 

Cryoultramicrotomy is an essential tool for effective polymer 

sample preparation for atomic force microscopy (AFM) 

in order to get rid of the polymer skin from processing and 

to ensure a smooth surface for analysis. We present an 

AFM study of the effect of tensile stress on a cryotomed 

impact copolymer (ICP), a multicomponent material typically 

used in automotive and appliance applications where a 

balance of stiffness and toughness is needed to investigate 

material deformation and interface adhesion as a function 

of tensile stress. 

Article 

The fi eld of atomic force microscopy (AFM), which was 

invented in the mid 1980’s, has revolutionized our capabilities 

to explore and understand nanoscale phenomena by 

allowing unprecedented characterization of surface and 

interface reactions and molecular and sub-molecular structures. 

Especially with commercial instruments available for 

widespread academic and industrial research beginning in 

the early 1990’s, the atomic force microscope (AFM) has 

become a main tool in the suite of techniques available for 

characterization, and is included in most characterization 

facilities alongside optical and electron microscopes. 

AFM now can routinely provide ~10nm lateral resolution 

and angstrom vertical resolution on a variety of surfaces 

and in fl exible environments including ambient and in situ 

fl uid imaging and is routinely used to provide a wealth 

of information including topography, mechanical properties, 

electrical and magnetic properties on a variety of 

materials ranging from biological cells to semiconductors 

to polymers. The heart of the AFM measurements lies in 

the precisely monitored interaction between a very sharp 

tip (~10nm in diameter) mounted on a cantilever (typically 

100’s of microns long, tens of microns wide and a few 

microns thick) and the surface of interest via optical detection. 

Through this tip-sample interaction, multiple surface 

properties can be probed on the nanoscale, including 

nanomechanical properties of polymeric materials. Specifi 

cally, an AFM mode called tapping mode or amplitude 

modulation mode is employed to image polymeric surfaces 

where the cantilever is oscillated at a resonance frequency 

and thus gently “taps” along the surface through intermittent 

contact, resolving features in the material based on its 

mechanical properties such as stiffness and other viscoelastic 

properties. 

Sample preparation of polymeric samples via ultracryomicrotomy 

for AFM analysis is critical for two reasons. First, 

a smooth surface is essential for effective AFM analysis 

as the maximum vertical range on AFM is typically less 

than 5 um. This is also means that if there are features that 

are taller than 5 um (or whatever the specifi cation on the 

particular AFM instrument), that surface will not be able 

to be imaged. Second, many polymer materials come in a 

processed form where the material has either been injection 

molded or compression molded and thus forms a rough 

skin on the surface that is not representative of the bulk 

material and needs to be removed. 

Cryomicrotomy is able to remove the surface skin and provide 

a smooth surface (all at cold temperatures below Tg 

of the polymer, which is essential, otherwise the surface 

features of interest will not be resolved). The Leica EM UC6 

system provides convenient AFM attachments where samples 

can be cryofaced in conjunction with the Leica EM FC6 

and directly transferred to the AFM for convenient analysis 

without removing the specimen, ensuring a smooth fl at 

surface for AFM imaging. 

We show here AFM data examining the rubber/matrix 

interface of a commercially important material, a polypropylene 

based impact copolymer. The interface of the two 

components in this material is examined by inserting a 

cryotomed dogbone of the impact copolymer material into 

an AFM tensile stage (NanoRack Asylum Research). 

Dalia G. Yablon 

NANOTECHNOLOGY 17

INDUSTRY 

Fig. 1 a: Stretching of impact Copolymer in neutral position Fig. 1 b: Stretching of impact Copolymer in stretched position 

18 reSOLUTION 

Fig. 1 c: Stretching of impact Copolymer in 

overnight after stretching 

A tensile stress is then exerted on the dogbone and the 

material imaged under tensile stress with AFM (MFP-3D 

Asylum Research). The dogbone was cryotomed in the Leica 

EM UC6 / FC6 system with a custom-built hemispherical 

accessory onto which the dogbone is glued allowing for 

the dogbone mid section to protrude out towards the knife. 

Cryotoming conditions were at -60C, a diamond knife 

speed of 0.3-0.6 mm/s and feed of 100-250 nm. Shown 

in Figure 1a is an AFM tapping mode phase image with 

approximately dimensions of 5 um x 5 um of a cryotomed 

impact copolymer. The commercial impact copolymer used 

for this study is composed of a polypropylene (PP) matrix 

with micron-sized domains of ethylene-propylene (EP) rubber 

domains, which further contain ethylene inclusions 

produced in a serial polymerization reactor. In the image, 

the PP matrix is observed as the surrounding purple medium, 

and the EP rubber domain as the large round bright 

yellow domain in the middle. Within this EP rubber domain 

there is a further smaller purple inclusion which is composed 

of ethylene. The color contrast in this phase image 

is due to a convolution of various mechanical properties 

where the EP rubber is softer than the surrounding stiffer 

PP matrix.

Fig. 2 a,b: Large scale image of a topography... and b: phase of a crack in polypropylene matrix 

Fig. 3 a,b: High resolution image of a topography... and b: of a crack in polypropylene matrix 

In Figure 1b, this impact copolymer was elongated in 

the direction of the black arrow by 1.7% (this elongation 

length is below the yield strain of PP) resulting in the AFM 

image shown in Figure 1b where the same rubber component 

is tracked from Figure 1a. Several new features are 

visible in this new image. First, rips and tears that were 

present within the rubber in Figure 1a (circled in blue), 

have now grown and elongated in Figure 1b (also circled 

in blue). Second, stretch marks (circled in red) between 

the rubber and the polypropylene matrix have developed 

at the north and south poles of the rubber domain, indicating 

a mismatch in Poisson ratios between the EP and PP 

materials. If the EP domain is stretched mainly along the 

equatorial line as in the experiment conducted here, then 

stretch marks would develop mainly at top and bottom of 

the EP rubber domain as observed in the AFM image in 

Figure 1b. 

INDUSTRY 

NANOTECHNOLOGY 19

INDUSTRY 

20 reSOLUTION 

Furthermore these marks are asymmetric about the EP rubber 

domain and appear to be most prominent at the bottom 

of the domain, though stretch marks are also observed on 

the top portion of the domain. The sample was allowed 

to sit overnight at 1.7% elongation and the next morning 

revealed a disappearance of the stretch marks as shown in 

the AFM image of Figure 1c, suggesting the yielding of the 

PP matrix overnight. 

Finally, the effect of the stress within the PP matrix at 2% 

elongation is shown in Figure 2. Both topography (a) and 

phase (b) images of a large-area (15um) scan size show a 

number of areas where cracks have formed at the EP-PP 

interface and propagated into the PP matrix; some of the 

cracks are highlighted in blue/orange circles. The developments 

of cracks or shear bands and micro-voids may come 

from stress amplifi cation in the ICP material due to the presence 

of EP rubber domains. Maximum stress amplifi cation 

by a spherical EP rubber domain is inversely proportional 

to the square root of the crack tip radius and occurs at the 

poles of the EP rubber domain. All these cracks and shear 

bands in Figure 2 appear to originate at the polar locations 

of the EP rubber domains, probably at sharp corners of the 

rubber domain with extremely small crack tip radii (and 

therefore maximum stress amplifi cation resulting in a stress 

singularity at that point). The appearance of these shear 

bands and micro-voids suggests that the local stresses well 

exceed the yield stress despite the 2% global deformation. 

The larger cracks propagate several microns within the 

polypropylene matrix. However, there are also several 

cracks with signifi cantly smaller dimensions of a couple 

hundred nm in length and tens of nm in width. Zooming in 

on the crack circled in orange from Figure 2 is shown in 

Figure 3 and reveals tiny PP fi brils stretching across the 

entire width of the track, as shown in the corresponding 

topography 3(a) and phase 3(b) images, at about 45 degree 

to the stretching direction suggesting that the cracking is 

induced by shear deformation. This particular crack is measured 

to be ~80 nm in depth and ~600 nm in width. 

Summary 

Morphology and interface adhesion of an impact copolymer 

(ICP) were studied using atomic force microscopy. Effects 

of deformation were observed within both PP and EP 

components as well as at the interface between the two 

materials. A continued stretching of the ICP could lead to 

delamination of EP from PP matrix. The strain required to 

separate the EP domains from the PP matrix could be used 

as a measure of the interfacial adhesion between EP and 

PP. Most importantly, the corresponding local interfacial 

stretching extent or void length between EP and PP upon 

delamination, which can be measured directly by AFM, can 

be used to calculate the interfacial strength between EP 

and PP. Presently, there are no direct measurement methods 

available to determine interfacial adhesive strength 

of nano- and microscale domains within polymer blends, 

especially blends generated in situ in polymerization reactors. 

This AFM examination of micro-domain deformation 

described qualitatively here could be used for direct determination 

of interfacial adhesion in complex polymer containing 

materials such as blends and composites. 

Figures reproduction permission 

Figures 1, 2, and 3 are reprinted with permission from 

Microscopy and Analysis 25(3):11-13 (AM), 2011, Copyright 

2011 John Wiley and Sons Ltd. 

Contact 

Dr. Dalia G. Yablon 

Corporate Strategic Research 

ExxonMobil Research and Engineering 

Annandale, NJ 

dalia.g.yablon@exxonmobil.com 


Leica EM UC6 and Leica EM FC6 the predecessor models of Leica EM UC7 and Leica EM FC7 

Leica EM UC7 

Ultramicrotome for Perfect Sectioning at Room Temperature and Cryo 

Leica EM UC7 with Cryochamber EM FC7

University of Wollongong 

Electron Microscopy Centre 

Darren Attard1 and Tony Romeo2 1 Institute for Superconducting and Electronic Materials (ISEM), 

Australian Institute for Innovative Materials (AIIM) Facility, University 

of Wollongong, Squires Way, North Wollongong NSW 2500 

2 Intelligent Polymer Research Institute (IPRI), AIIM Facility, 

University of Wollongong, Squires Way, North Wollongong 

The University of Wollongong has a diverse range of materials 

research programs that includes metallurgy for mining, 

manufacturing, steel making and transport; polymers 

for solar cells, energy storage and bionic implants; and 

superconducting and electronic materials for commercialization, 

energy storage, telecommunications and medical 

applications. Electron microscopy is an integral part of this 

research, due to the chemical and structural information 

that can be provided down to the atomic scale. However, 

as the performance of electron microscopes and analytical 

techniques has evolved, they have become increasingly 

sensitive to environmental effects and the quality of specimen 

preparation has become even more of a crucial factor 

to the success of applied materials studies. 

To address a number of performance issues related to 

electron microscopy, the University of Wollongong has 

constructed a purpose built electron microscopy centre 

that was completed in July 2011, and purchased a comprehensive 

suite of specimen preparation equipment that 

is now operational for materials and life sciences. The 

centre will house up to 7 electron microscopes with two 

preparation laboratories, and has been designed to exceed 

the environmental specifi cations for the current generation 

of commercially available electron microscopes including 

aberration corrected S/TEMs. Currently two FEGSEm’s and 

a TEM have been relocated to the facility and are fully operational. 

Leica was selected as a major supplier for the 

specimen preparation equipment based on equipment 

specifi cations, product integration, demonstrated capability, 

ease of use, automation and applications support. These 

will be supplemented with an Leica M205 A stereo, Leica 

DM2500 M and Leica DM6000 M optical microscopes for 

quality control during specimen preparation and their value 

as stand-alone research tools. These factors will facilitate 

production of high quality specimens, and the training of a 

large user-base typical of a university environment. 

The Leica EM TXP, EM TIC020 and EM RES101 were purchased 

to prepare materials for SEM and TEM based studies. 

The Leica EM TXP was selected for its ability to prepare 

TEM specimens and perform target grinding. The ability to 

perform these functions while being viewed directly to assess 

quality and increase specimen throughput was seen 

as a major advantage. The Leica EM TIC020 was selected 

for its ability to prepare large, damage free areas of specimens 

for low voltage, high resolution SEM imaging, Electron 

Backscatter Diffraction (EBSD) and Energy Dispersive 

Spectroscopy (EDS). The Leica EM TXP and EM TIC020 instruments 

have already been utilized for the preparation of 

Magnesium diboride (MgB 2 ) superconducting wires. MgB 2 

is a challenge to prepare because it is brittle and sensitive 

to water. Of particular interest are defects such as pores, 

cracks or impurities at the Mg and B grain boundaries after 

processing, which preparation artefacts can infl uence. 

Transverse and longitudinal orientations of MgB 2 wires encapsulated 

in Niobium (Nb) metal and Inconel alloys, were 

prepared using the Leica EM TXP and EM TIC020 instruments. 

The specimens were glued to the small Leica EM 

TIC sample holders, then clamped for preparation on the 

Leica EM TXP using the procedure in Table 1. 

Tool Insert RPM Force limit Pump/mL E-W speed/mms-1 Diamond disc cutter 15000 - 18 0.025 

Silicon carbide foil, 15μm 1000 8 8 0.2 

Diamond lapping foil, 9μm 2000 8 8 0.2 

Table 1: Leica EM TXP preparation of MgB 2 superconduting wires 

INDUSTRY 

Darren Attard Tony Romeo 

NANOTECHNOLOGY 21

INDUSTRY 

Fig. 1: a Ion polished transverse section of the superconducting wire, and b detail of the MgB 2 core. 

Fig. 2: a Ion polished longitudinal section of the superconducting wire, and b detail of the MgB 2 core. 

Fig. 3: X-ray maps of the MgB2 superconducting material showing a the distribution of a) Magnesium, b) Boron, c) oxygen and 

d) all elements overlaid to show the spatial relationship. 

22 reSOLUTION 

Ethanol was used as a lubricant and dispensed using the 

Leica EM TXP pump. The polishing procedure on the Leica 

EM TXP only used coarse lapping foils to provide a plane 

surface for fi nal preparation with the Leica EM TIC020. A 

mechanical polishing procedure will be developed shortly 

to produce the best possible surface fi nish. 

To minimize the effects of surface oxidation after preparation 

on the Leica EM TXP, the specimens were transferred 

directly to the Leica EM TIC020 and placed under vacuum. 

The specimens were ion milled with gun energies of 7kV 

and gun currents of 2.6mA. After polishing, the specimens 

were transferred directly to the SEM to minimize exposure 

to air. Figure 1a shows the transverse section with 

the Nb metal sheath encapsulating the MgB 2 superconducting 

core. The ion polished surface and the interface 

between the dissimilar materials, are free of polishing 

artefacts. Figure 1b shows the MgB 2 core in more detail, 

with pores present at some grain boundaries. Figure 2a 

shows the longitudinal section of the superconducting 

wire where the elongated grains in the drawing direction 

are easily observed. Figure 2b shows the MgB2 core in 

more detail, where no physical defects were observed. 

Figure 3 shows X-ray maps of a) Mg, b) B and c) O in the 

transverse section of the MgB 2 superconducting wire. The 

x-ray maps show the elemental concentration as a function 

of the colour intensity in each respective map. Figure 

3d shows all of the elemental maps superimposed. 

Here, it can be seen that O is present in low concentrations, 

mostly at the Mg grain boundaries where the superconducting 

properties can be affected. 

The electron microscopy centre will also need to support 

growing areas in biology-based research projects needing 

TEM and SEM. A Leica EM UC7 ultramicrotome and EM 

FC7 cryo-attachment have been purchased to section soft 

polymers and biological materials to cater for the new 

research interests. Similarly, a Leica EM CPD030 critical 

point dryer will allow controlled drying of soft biological 

tissues and gels for SEM observation, and completes the 

suite of Leica instruments that will be installed in the new 

centre. 

Contact 

Darren Attard 

Microscopist and Services Coordinator 

AIIM Electron Microscopy Centre, Innovation Campus 

University of Wollongong, Sydney 

darrena@uow.edu.au


Leica EM TXP and Leica EM TIC020 (the predecessor of Leica EM TIC 3X) 

Leica EM TXP 

Target Surfacing System 

Leica EM TIC 3X 

Ion Beam Slope Cutter 

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Order no.: English 11924257 ∙ IX/12/LX/B.H. ∙ Copyright © by Leica Mikrosysteme GmbH, 

Vienna, Austria, 2012. Subject to modifications. LEICA and the Leica Logo are registered 

trademarks of Leica Microsystems IR GmbH. 

IMPRINT 

Publisher 

Leica Mikrosysteme GmbH 

Hernalser Hauptstraße 219 

A - 1170 Vienna, Austria 

www.leica-microsystems.com 

Editor in Chief 

Ian Lamswood 

Marketing Manager LNT 

Leica Microsystems, Vienna, Austria 

Ian.Lamswood@leica-microsystems.com 

Contributing Editors 

Akos Kulik 

Cathy Johnson 

Dalia G. Yablon 

Daniel Studer 

Daniel Althof 

Darren Attard 

Günter P. Resch 

Jeff P. Gorski 

Karin Trimmel 

Marlene Brandstetter 

Martin W. Goldberg 

Nicole Fellner 

Thérèse Hillmann 

Tony Romeo 

Layout 

Uwe Neumann, 

Leica Microsystems GmbH 

Communications & Corporate Identity 

Isabelle Elmann 

Marketing LNT 

Leica Microsystems, Vienna, Austria 

Cover Picture 

Cathy Johnson 

Leica Microsystems 

Printing Date 

4th October 2012 

INDUSTRY 

NANOTECHNOLOGY 23

Image Is Everything 

And We‘ve Got It Covered 

The new Coater range from Leica

reSolution_LNT_No1_en - Leica Microsystems

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