CUSTOMER MAGAZINE

FOR NANOTECHNOLOGY

reSOLUTIONNo.

01

, A

utu

mn

20

12

2 reSOLUTION

B I O L O G Y

Capturing neuro transmitter receptors 04

and ion channels:

High-resolution techniques to localizemembrane 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 applicationsfor 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

CON

TEN

T 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

EDIT

OR

IAL 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!Best Wishes,

Ian LamswoodMarketing Manager

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 andTechnology

2 Smithsonian InstitutionSource: 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

ddddding!es,

wood

NANOTECHNOLOGY 3

B I O L O G Y

Capturing Neurotransmitter Receptors and Ion Channels

High-resolution Techniquesto Localize Membrane ProteinsDaniel Althof1,2, Akos Kulik1,3

1Department of Anatomy and Cell Biology, Institute of Neuroanatomy2Spemann Graduate School of Biology and Medicine, University of Freiburg, Germany3Department 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 ele-ments of neurons. The impact of the activation of these proteins on synaptic integration and regulation of transmit-ter release depends on their precise location relative to synapses, as well as on the density and coupling of mol-ecules in microcompartments of the cells. High-resolution qualitative and quantitative visualization of membrane-bound 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 neu-rons 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 synap-tic membrane, contains a large variety of molecules such as receptors, ion channels, and associated structural pro-teins, whose precise subcellular organization facilitates its proper function. A large number of studies have provided evidence that the location of these proteins and their posi-tion relative to synapses substantially affects their func-tional roles. Neurotransmitter receptors, localized to the synaptic membrane of postsynaptic compartments of neu-rons, 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 con-trast, receptors localized to the extrasynaptic plasma mem-brane, remote from synaptic sites, are activated by spilled-over 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 scale1. 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 po-tential.

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 neu-rotransmitter release, thereby playing a role in the presyn-aptic 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 cells1.

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 lo-cation relative to excitatory and inhib-itory 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 ad-vanced high-resolution immunocyto-chemical methods have been widely used: (i) preembedding immunogold, (ii) postembedding immunogold, and (iii) sodium dodecyl sulfate (SDS)-di-gested 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

Akos Kulik Daniel Althof

Fig. 1

4 reSOLUTION

B I O L O G Y

in order to facilitate proper penetra-tion. 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 re-action and the location of the protein at extra- and perisynaptic sites can be determined. Synaptic proteins, how-ever, cannot be detected using this method, most likely due to the inac-cessibility of the epitopes in the syn-aptic specializations of fi xed tissues2. (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 technique2. (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 ma-chine (Leica EM BAF060). Proteins are allocated to either the protoplasmic faces (P-faces) or the exoplasmic faces (E-faces) of plasma membranes. Molecules are immo-bilized 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 deposit3. 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, be-cause membrane proteins are exposed on the two-dimen-sional 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 morphologi-cal structures is diffi cult, therefore, it is necessary to use marker proteins to facilitate the identifi cation of fractured membranes6. Second, the separation of membrane proteins

to P-face or E-face is unpredictable: some proteins are pref-erentially allocated to either the P-face, such as GABA (B1), Kir3.24 or the E-face, such as AMPA receptors5, whereas others, like gluRδ26 are localized to both faces. Thus, for quantitative studies, the allocation of the molecules should carefully be examined7.

Taken together, these three immunocytochemical tech-niques provide complementary information about the cel-lular and subcellular distribution of proteins and are widely used for high-resolution qualitative and quantitative analy-sis of receptor and ion channel localization and colocaliza-tion in post- and presynaptic compartments of neurons.

ContactDaniel Althof, Akos KulikDepartment of Anatomy and Cell Biology, Institute of Neuroanatomy, Spemann Graduate School of Biology and Medicine, University of Freiburg, GermanyDepartment of Physiology II, University of Freiburg, Germanyakos.kulik@physiologie.uni-freiburg.de

Fig. 1 & 2: Distribution and colocalization of GABA (B1) and Kir3.2 in dendrites of hippocam-pal cells as revealed by the SDS-digested freeze-fracture replica labeling technique. A, Immu-noparticles 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

Fig. 2

B I O L O G Y

NANOTECHNOLOGY 5

Leica EM HPM100High Pressure Freezer for Cryofi xationof Biological and Industrial Samples

References1. 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., Shige-moto, R. Compartment-dependent colocalization of Kir3.2-containing K+ channels and GABAB receptors in hippocampal pyramidal cells. J Neuroscience 26(16), 4289-4297. (2006)

5. Masugi-Tokita, M., Tarusawa, E., Watanabe, M., Molnar, E., Fujimoto, K., Shigemoto, R. Number and density of AMPA re-ceptors in individual synapses in the rat cerebellum as revealed by SDS-digested freeze-fracture replica labeling. J Neurosci-ence 27(8), 2135-2144. (2007a)

6. Masugi-Tokita, M., Shigemoto, R. High-resolution quantita-tive 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 com-bined 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 BAF060Freeze Fracture System

6 reSOLUTION

Biological ElectronMicroscopy

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 mi-

croscopes, TIRF microscopy and live cell imaging) as well as an ultra high resolution (<0.5nm) fi eld emission in-lens scanning

electron microscope (Hitachi S5200) and a routine transmission EM with high tilt stage for tomography (Hitachi H7600).

These instruments are supported by a comprehensive range of sample preparation equipment including a Leica EM PACT high pressure freezer, Leica metal mirror and plunge freezers (MM80 and CPC), Leica EM AFS freeze substitu-tion instrument, Leica cryo-ultramicrotome, Baltech (now Leica) CPD030 critical point drier, Cressington vacuum units for carbon evaporation and for ultra high resolution metal coating (e.g. chromium). Our aim is to develop the best possible methods for examining and analysing biological ultrastructure both in thin sections (TEM) and on biologi-cal surfaces (SEM), and then to offer these techniques to internal and external clients.

Most importantly, methods are generally developed as part of intensively EM oriented internal research projects. We work with a wide range of samples including plant tis-sues, single molecules, polymers, all types of animal tis-sue and culture cells, which may be processed for surface imaging, thin sectioning and immuno-gold labelling. One particular research interest is to understand how nuclear pore complexes (NPCs) carry out the controlled transport of molecules such as proteins and RNAs to and from the cell nucleus. We look at NPC structure using SEM by isolating the nuclear envelope (which encloses the nucleus and con-tains the NPCs), critical point drying them and then coating with a 1.5 nm thick fi lm of chromium (Fig.1). Such samples can be labelled with gold-tagged antibodies to determine the position of specifi c proteins (Fig.2).

To look at how molecules travel through the NPC we have to look at cross sections in the TEM. We use baker’s yeast because in this organism it is easy to genetically manipulate the proteins that make up the NPC. We can then discover how those proteins are involved in NPC structure and func-tion. The problem however is that yeast has a cell wall which is resistant to chemical fi xation. Transport through the NPC is also extremely rapid, so molecules are rarely caught in tran-sit. Both these problems are solved by high pressure freezing followed by low temperature fi xation and embedding (Fig.3).

B I O L O G Y

Fig. 1: Field emission scanning electron micrograph of an isolated Xenopus laevis oocyte nuclear envelope showing the outer surface the outer nuclear membrane and the nuclear pore complexes.

Fig. 2: Field emission scanning electron micrograph of an isolated Xenopus laevis oocyte nuclear envelope showing the inner facethe inner nuclear membrane and the nuclear pore complex baskets immuno-gold labelled for a NPC protein. The secondary electron image showing the sample structure is in red and the backscatter image which detects the position of the gold particle is in yellow.

Dr. Martin W. Goldberg, Durham University, UK

NANOTECHNOLOGY 7

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).

ContactDr. Martin W. Goldberg,Durham University, UKm.w.goldberg@durham.ac.uk

B I O L O G Y

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 ACE200Low Vacuum Coater

Instruments related to this sample preparation:

Leica EM TPAutomated Routine Tissue Processor

Leica EM ACE600High Vacuum Coater

Leica EM CPD300Automated Critical Point Dryer

8 reSOLUTION

Perusing alternatives for automated staining of TEM thin sections

Substitutes for Uranyl Acetate in TEM Thin Section Post-StainingNicole 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, Austria2University of Applied Sciences, Wiener Neustadt, Austria3Campus Science Support Facilities GmbH, Vienna, Austria

B I O L O G Y

Introduction

Contrast in transmission electron mi-croscopy (TEM) is mainly produced by electron scattering at the speci-men: 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, bio-logical samples show hardly any in-herent 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 ele-ments 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 shad-owing). 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 ap-proach, 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 satu-rated solution in the range of 3.5 to 4.0. The addition of alcohol, especially methanol, increases the solubil-ity (Hayat, 2000). Uranyl acetate strongly stains pro-teins 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 eithermanually or automatically, both techniques have their ad-vantages. For manual staining, a grid is fl oated, section-side 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 auto-mated 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 re-quires 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 pre-pared solutions are available, handling of solid UA can be avoided. Two reagents described in literature as replace-ments 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 phos-phate buffer and post-fi xed with 2.0% osmium tetroxide.

Nicole Fellner (left), Marlene Brandstetter (in the middle of the front row), Günter Resch (in the middle of the back row), Harald Kotisch (right)

NANOTECHNOLOGY 9

B I O L O G Y

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 pep-tide 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

2O was used for further experi-

ments, 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 con-trast 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 prob-lem 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 cis-dichlordiamineplatinum (II) with thymidine (Inaga et al., 2007). The reagent can be ordered directly from the Japa-nese producer Nisshin (http://nisshin-em.co.jp). Pt Blue is a hazardous material, which may cause eye irritation, can-cer 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 wa-ter, 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 qual-ity to pictures taken from sections stained with UA. Differ-ences can be found in a more intensely stained mitochon-drial 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 ddH2O Lead citrate 5 min 2 min ddH

2O

OTE 25 min 6 min ddH2O Lead citrate 5 min 2 min ddH

2O

Pt Blue 10 min 2 min ddH2O Lead citrate 5 min 2 min ddH

2O

Table 1: Details of manual staining procedures

Stain 1 Time Washing Stain 2 Time Washing

UA 30 min 2 min 20 sec ddH2O Lead citrate 7 min 2 min 20 sec ddH

2O

OTE 40 min 5 min ddH2O Lead citrate 7 min 2 min ddH

2O

Pt Blue 30 min 2 min 20 sec ddH2O Lead citrate 7 min 2 min 20 sec ddH

2O

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.

10 reSOLUTION

B I O L O G Y

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. Man-ual 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 sec-tion 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 interpre-tation 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. Further-more, 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 fur-ther 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

up to 20 grids at once and all steps are carried outautomatically, this becomes less of a disadvantage of OTE. From the economic point of view, OTE was by far the cheap-est 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 replace-ment 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 Microsys-tems, 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 Cel-lular Nanostructure.”

ContactDr. Günter ReschHead of Electron Microscopy, Campus Science Support Facilities GmbH, Vienna, Austriaguenter.resch@csf.ac.at

Instruments related to this sample preparation:

Leica EM TRIM2Specimen Trimming Device for TEM, SEM, LM

Leica EM AC20Automatic Constrasting Instrument for Ultrathin Sections

Leica EM UC7Ultramicrotome for Perfect Sectioning at Room Temperature and Cryo

NANOTECHNOLOGY 11

B I O L O G Y

References

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

html.

2. Dykstra, M.J. 1992. Biological Electron Microscopy: Ther-

ory, Techniques, and Troubleshooting. Plenum Press, New

York.

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

ning and Transmission Electron Microscopy: An Introduction.

W.H. Freeman and Company, New York.

4. Hayat, M.A. 2000. Principles and Techniques of Electron Mi-

croscopy: Biological Applications. 4th ed. Cambridge University

Press, Cambridge.

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

T., Nakane, H., Naguro, T. 2009. Low vacuum scanning elec-

tron 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 ac-

etate 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 Mi-

crosporidia (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 sec-

tions. 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., Ya-

mamoto, 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.

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.

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 tis-sue. (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 gly-cogen granules are clearly visible in this specimen stained with Pt Blue (1:200). Scale bar: 1 μm.

12 reSOLUTION

B I O L O G Y

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 CulturesJeff 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

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 com-plexes, termed biomineralization foci (BMF), in which the fi rst crystals of hydroxyapatite mineral are deposited (Mi-dura 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 plat-ing 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 diam-eter. However, the fi rst mineral crystals are not detected until 78 h after plating of UMR106-01 cells and are local-ized 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 cul-ture 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

occur. Thus, mineralization within BMF is a temporally synchronized process. However, understanding the bio-chemical 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 Ot-tensmeyer, 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 sec-tioning. Since most prior studies have not consistently avoided water during specimen processing, the true im-pact 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 non-aqueous 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 distri-butions in other cells such as muscle.

Jeff P. Gorski Daniel Studer

Thérèse Hillmann

NANOTECHNOLOGY 13

B I O L O G Y

Calcium and phosphorus can be lost from samples either after high pressure freezing and freeze-substitution or dur-ing 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 ap-pearance of the fi rst crystals of mineral within BMF (Mi-dura 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 con-clusions. 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 cul-tures 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 underly-ing detail which represents a range of irregularly shaped spherical bodies from about 50 to 800 nanometers in diam-eter (not shown). However, the presence of these “white” areas raised immediate concerns regarding the loss of inor-ganic or organic substances. In particular, when compared with similar BMF regions from paired, conventionally fi xed cultures (Fig. 2), we hypothesized that “white” spots rep-resented 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 conven-tionally 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 sub-stituted dry sectioning for wet sectioning of high pressure frozen, freeze substituted cultures. Calcium and phospho-rus retention is improved in dry sectioned high pressure fro-zen, 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 synchro-nous manner facilitates these direct comparisons among different cultures (Wang et al., 2009).In addition, electron spectroscopic imaging demonstrates that the dark appear-ing 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 (Dries-sens 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 fo-cal 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.

Fig. 1: The single and dual vesicle models of extracellular mineralization. Upper panel, single matrix vesicle model. Calcium and phosphorus ions are progressively concen-trated 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 (yel-low), nucleation of initial mineral crystals occurs leading to breakage of the vesicle and release of crystals which can propagate additional crystals within the surround-ing extracellular collagenous matrix. Lower panel, dual vesicle model. Calcium and phosphorus are progressively concentrated within two different populations of vesi-cles which biochemically display distinct functional distributions including Ca+2 pump-ing 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 propa-gate additional crystals within the surrounding extracellular collagenous matrix.

Leica EM PACT2High Pressure Freezer with Rapid Transfer System

Instrument related to this samplepreparation:

14 reSOLUTION

B I O L O G Y

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

NANOTECHNOLOGY 15

B I O L O G Y

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 en-richment 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 publica-tions. 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-Kan-sas City, 650 East 25th Street, Kansas City, MO 64108.Phone: 815-235-2537; fax: 816-235-5524; gorskij@umkc.edu

ContactJeff P. Gorski, Ph.D.Professor UMKC Center of Excellence in Mineralized TissuesDepartment of Oral BiologySchool of DentistryUniversity of Missouri-Kansas Citygorskij@umkc.eduDr. sc. nat. Daniel StuderInstitute of Anatomy, University of Bern, Bern, Switzerlanddaniel.studer@ana.unibe.ch

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.

References1. Anderson, H.C. (1967) Electron microscopic studies of indu-

ced cartilage development and calcifi cation. J Cell Biol 35: 81-101.

2. Arsenault, A.L., F.P. Ottensmeyer (1984) Visualization of ear-ly 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 ca-tion. J Ultrastruct Res 20, 33-50.

4. Driessens, F.C., J.W. van Dijk, J.M. Borggreven (1978) Bio-logical 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. Mi-dura (2004) Extracellular bone acidic glycoprotein-75 defi nes condensed mesenchyme regions to be mineralized and loca-lizes with bone sialoprotein during intramembranous bone formation. J Biol Chem 279: 25455–25463.

6. Huffman, N.T., J.A. Keightley, C. Chaoying, R.J. Midu-ra, 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 mine-ralization within biomineralization foci. J Biol Chem 282: 26002–26013.

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. Suk-hu, 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 Mineralizi-ng 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 micros-pectroscopy of biomineralization foci in UMR 106 osteoblas-tic cultures reveals temporally synchronized protein changes preceding and accompanying mineral crystal deposition. J Biol Chem 284: 7100–7113.

A Word on Cathodoluminescence

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 elec-

tronic 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.

CL data can indicate defects such as imperfections or impu-rities within the microstructure of a material phase. These defects can have an effect on the material’s optical, electri-cal 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 tech-nique in the characterization of materials. Image resolution is dependent on instrument confi guration, experimental pa-rameters 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 Instituteof Standards and Technology (NIST) in Gaithersburg, MD. This conference was sponsored by the Microbeam Analy-sis Society (MAS), and was co-sponsored by the AustralianMicrobeam Analysis Society (AMAS). The four day pro-gram included a pre-conference tutorial targeted for the CL novice on October 24th. The remaining three days included a combination of technical presentations, hands-on labora-tory demonstrations and a contributed poster session. Pre-sentation topics included: CL theory, data quantifi cation, advances in instrumentation, analysis and databases. Ap-plications 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.

ContactCathy JohnsonLeica Microsystems Nanotechnology Division1700 Leider LaneBuffalo Grove, IL 60089Cathy.Johnson@leica-microsystems.com

I N D U S T R Y

Cathy Johnson, Leica Microsystems

Instruments related to this sample preparation:

Leica EM TIC 3XIon Beam Slope Cutter

Leica EM TXPTarget Surfacing System

Leica EM RES101Ion Milling System

Instruments rel

Panchromatic cathodoluminescence image of cassiterite mineral (SnO2)

Image Credits

Scott Wight1, Ed Vicenzi2, Doug Meier1, and Kurt Benkstein1

1 National Institute of Standards and Technology2 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

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

NANOTECHNOLOGY 17

I N D U S T R Y

Atomic Force MicroscopyStudy of a Stretched ImpactCopolymerDalia 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 na-noscale lateral and sub-nanometer vertical resolution.Cryoultramicrotomy is an essential tool for effective poly-mer 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 typical-ly 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 capabili-ties to explore and understand nanoscale phenomena by allowing unprecedented characterization of surface and interface reactions and molecular and sub-molecular struc-tures. 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 proper-ties, 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 de-tection. Through this tip-sample interaction, multiple sur-face properties can be probed on the nanoscale, including

nanomechanical properties of polymeric materials. Spe-cifi 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 intermit-tent contact, resolving features in the material based on its mechanical properties such as stiffness and other visco-elastic properties.

Sample preparation of polymeric samples via ultracryomi-crotomy 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 injec-tion 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 pro-vide 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 sam-ples 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 polypro-pylene 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

18 reSOLUTION

I N D U S T R Y

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 Lei-ca EM UC6 / FC6 system with a custom-built hemispheri-cal 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) rub-ber domains, which further contain ethylene inclusions produced in a serial polymerization reactor. In the image, the PP matrix is observed as the surrounding purple me-dium, 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 com-posed 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. 1 a: Stretching of impact Copolymer in neutral position Fig. 1 b: Stretching of impact Copolymer in stretched position

Fig. 1 c: Stretching of impact Copolymer inovernight after stretching

NANOTECHNOLOGY 19

I N D U S T R Y

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 compo-nent 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, indica-ting 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.

20 reSOLUTION

I N D U S T R Y

Furthermore these marks are asymmetric about the EP rub-ber 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 develop-ments of cracks or shear bands and micro-voids may come from stress amplifi cation in the ICP material due to the pres-ence 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 mea-sured to be ~80 nm in depth and ~600 nm in width.

Summary

Morphology and interface adhesion of an impact copoly-mer (ICP) were studied using atomic force microscopy. Ef-fects 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 meth-ods available to determine interfacial adhesive strength of nano- and microscale domains within polymer blends, especially blends generated in situ in polymerization reac-tors. This AFM examination of micro-domain deformation described qualitatively here could be used for direct deter-mination of interfacial adhesion in complex polymer con-taining materials such as blends and composites.

Figures reproduction permissionFigures 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.

ContactDr. Dalia G. YablonCorporate Strategic ResearchExxonMobil Research and EngineeringAnnandale, NJdalia.g.yablon@exxonmobil.com

Instruments related to this sample preparation:

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

Leica EM UC7 with Cryochamber EM FC7Leica EM UC7Ultramicrotome for Perfect Sectioning at Room Temperature and Cryo

NANOTECHNOLOGY 21

I N D U S T R Y

University of WollongongElectron Microscopy CentreDarren 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 25002 Intelligent Polymer Research Institute (IPRI), AIIM Facility,

University of Wollongong, Squires Way, North Wollongong

The University of Wollongong has a diverse range of ma-terials research programs that includes metallurgy for min-ing, manufacturing, steel making and transport; polymers for solar cells, energy storage and bionic implants; and superconducting and electronic materials for commercial-ization, 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 analyti-cal techniques has evolved, they have become increasingly sensitive to environmental effects and the quality of speci-men 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 com-prehensive 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 op-erational. Leica was selected as a major supplier for the specimen preparation equipment based on equipment specifi cations, product integration, demonstrated capabili-ty, 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 pur-chased to prepare materials for SEM and TEM based stud-ies. 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 as-sess 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 speci-mens for low voltage, high resolution SEM imaging, Elec-tron Backscatter Diffraction (EBSD) and Energy Dispersive Spectroscopy (EDS). The Leica EM TXP and EM TIC020 in-struments 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 af-ter processing, which preparation artefacts can infl uence. Transverse and longitudinal orientations of MgB

2 wires en-

capsulated in Niobium (Nb) metal and Inconel alloys, were prepared using the Leica EM TXP and EM TIC020 instru-ments. 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.

Darren Attard Tony Romeo

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 MgB2 superconduting wires

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 prepara-tion 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 expo-sure to air. Figure 1a shows the transverse section with the Nb metal sheath encapsulating the MgB

2 supercon-

ducting 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 func-tion of the colour intensity in each respective map. Fig-ure 3d shows all of the elemental maps superimposed. Here, it can be seen that O is present in low concentra-tions, mostly at the Mg grain boundaries where the su-perconducting 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.

ContactDarren AttardMicroscopist and Services CoordinatorAIIM Electron Microscopy Centre, Innovation CampusUniversity of Wollongong, Sydneydarrena@uow.edu.au

Fig. 1: a Ion polished transverse section of the superconducting wire, and b detail of the MgB2 core.

Fig. 2: a Ion polished longitudinal section of the superconducting wire, and b detail of the MgB2 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.

I N D U S T R Y

IMPRINT

PublisherLeica Mikrosysteme GmbHHernalser Hauptstraße 219A - 1170 Vienna, Austriawww.leica-microsystems.com

Editor in ChiefIan LamswoodMarketing Manager LNTLeica Microsystems, Vienna, AustriaIan.Lamswood@leica-microsystems.com

Contributing EditorsAkos KulikCathy JohnsonDalia G. YablonDaniel StuderDaniel AlthofDarren AttardGünter P. ReschJeff P. GorskiKarin TrimmelMarlene BrandstetterMartin W. GoldbergNicole FellnerThérèse HillmannTony Romeo

LayoutUwe Neumann,Leica Microsystems GmbHCommunications & Corporate Identity

Isabelle ElmannMarketing LNTLeica Microsystems, Vienna, Austria

Cover PictureCathy JohnsonLeica Microsystems

Printing Date4th October 2012

NANOTECHNOLOGY 23

Register for UpcomingreSOLUTION IssuesWould you like to read future issues of reSOLUTION, too?The following link will take you straight to the registration form:

www.leica-microsystems.com/registration.01

Choose between different reSOLUTION editions and we would also be glad to hear your opinion of our magazine and any suggestions for topics.

You can find this issue and all past issues of reSOLUTION on our website at:

www.leica-microsystems.com/magazines.01

Instruments related to this sample preparation:

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

Leica EM TIC 3XIon Beam Slope Cutter

Leica EM TXPTarget Surfacing System

I N D U S T R Y

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.

Image Is Everything

And We‘ve Got It Covered

The new Coater range from Leica