TRANSMISSION ELECTRON MICROSCOPY

T RANSMISSION E LECTRON M ICROSCOPY F90 Fortgeschrittenen-Praktikum Universität Heidelberg Prof. Dr. Rasmus R. Schröder Anne Katrin Kast Last Update...
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T RANSMISSION E LECTRON M ICROSCOPY

F90 Fortgeschrittenen-Praktikum Universität Heidelberg

Prof. Dr. Rasmus R. Schröder Anne Katrin Kast Last Update: April 2016

Contents 1

Introduction

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Outline of Experiments 2.1 Day 1 . . . . . . . 2.2 Day 2 . . . . . . . 2.3 Day 3 . . . . . . . 2.4 Questionnaire . . .

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Expected Documentation of the Experiments 3.1 Day 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Day 2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Day 3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Experimental Procedure 4.1 Microscope Alignment – The EM109EL . . . . . . . . . . . . . . . 4.1.1 Turning the Microscope on . . . . . . . . . . . . . . . . . 4.1.2 The Control Panel . . . . . . . . . . . . . . . . . . . . . . 4.1.3 Beam Alignment . . . . . . . . . . . . . . . . . . . . . . . 4.1.4 Camera Flat-Field Correction . . . . . . . . . . . . . . . . 4.1.5 Inserting a Sample . . . . . . . . . . . . . . . . . . . . . . 4.1.6 Adjusting the Objective Astigmatism – Carbon Film Sample 4.2 Electron Diffraction . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Magnesium Oxide . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Calibration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Catalase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Helpful Features of Fiji 5.1 Opening images . . . . 5.2 Fast Fourier Transforms 5.3 Image Stacks . . . . . . 5.4 Scale Bars . . . . . . . .

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Provided Samples

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References / Suggested Reading

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Appendix

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1

Introduction

Transmission electron microscopy has in recent years developed into a – somewhat – versatile tool for physical as well as biomedical sciences. The basic interaction between electrons and sample, i.e. the interaction with the atoms in the sample consisting of atomic nuclei and electrons, is well understood and can be described by theories for the elastic electron-nucleus, and the inelastic electron-electron interaction (cf suggested literature, e.g. Williams and Carter [1], Reimer and Kohl [2], R. Erni [3], J. Rodenburg [4]...). However, necessary instrumentation for high-resolution, atomic imaging in a transmission electron microscope (TEM) had to be developed in a long process – culminating in the correction of lens aberrations realized in the last two decades. The correction of lens aberrations such as spherical aberration, coma, and more recently also chromatic aberration has allowed scientists to directly visualize columns of atoms in crystalline material and also individual carbon atoms in graphene (M. Haider et al. [5]). Figures 1.1 and 1.2 were adopted from the SALVE web-page (http://www.salveproject.de) [6], a recent collaborative project for optimizing imaging and electron spectroscopy on carbon-based materials. In materials science today, the TEM is used for high-resolution imaging as illustrated here, but also for studies based on more conventional electron scattering experiments, i.e. electron diffraction on crystalline material, as well as for analytical studies. Such studies utilize the special nature of inelastic interactions between the incident electrons and the electrons in the sample. In general, all these techniques are employed to obtain information about the structure of a material and its localized electronic properties. Imaging resolution is very often in the 1 A˚ to even sub-Å range. The resolution of the analytical signal is normally reduced to the typical delocalization of the inelastic interaction in the sample. In biomedical electron microscopy, the situation is quite similar: With advances in sample preparation and, in particular, constantly improving electron imaging detectors, it is now possible to image biomolecules such as e.g. proteins or protein complexes in their 3D structure at a quasi molecular level, sometimes in the range of 3 − 5 A˚ – more ˚ Figure 1.3 shows a typical image of such a «6 A˚ routinely in the range of 6 − 8 A. study» of the actin-myosin model complex, i.e. the force producing protein complex, which works in our muscles. If one compares the images of a materials sample with the images of the actin-

Figure 1.1: Strontium titanate as a typical materials science sample where in the past it was not possible to image individual columns in an oriented and thinned specimen. (A) shows the known atomic structure, which has been imaged in the past in conventional microscopes (B). Such images could only be interpreted by advanced simulation of image contrast from known models. (C) and (D) show images and more modern simulations of aberration corrected imaging. Figure adopted from SALVE web-page[6]. 1

Figure 1.2: Examples of conventional (upper row) and aberration corrected images (lower row) of a mixed crystalline/amorphous Si specimen. The direct imaging in the aberration corrected microscope shows the individual Si atom columns of the crystalline area (left part of the picture). This cannot be resolved in the conventional image. Figure adopted from SALVE web-page[6]. myosin complex one clearly realizes, that the level of resolution and atomic detail is completely different. The reason for this is very simple: The electrons used for imaging also damage the sample. This is obvious from the physics of inelastic interactions, their energy transfer and the fact, that inelastic interactions are more probable than elastic interactions for light material. For large Z materials (Z > 26), elastic interactions start to win. Biomedical samples, which are prepared in a glassy, vitrified aqueous buffer layer of about 50 − 100 nm thickness, are simply «boiled away» by our electron beams. Thus, images have to be taken at lower magnification and lower electron dose, which decreases resolution and increases noise. What will you learn from this experiment? As you might expect, TEMs which deliver sub-Å resolution or allow the imaging of a frozen sample at liquid nitrogen temperature are very specialized, expensive, and most of the time used for our research. A microscope – as small as the one used here in our practical course – will not have all this expensive instrumentation, but will get you acquainted with some very important physical facts, which we in our daily research still have to be aware of. Familiar topics in this respect are the optimal optical alignment of an imaging system in particle optics, basic imaging properties when working with a particle beam, the recording of an image and basic image processing steps. Also, electron diffraction experiments can be performed with our microscope, and very fundamental experiments to test the coherence of an electron beam and how it can be influenced by different beam forming parameters. In the end you should have a first feeling - and also the physical basis for it . . . - what it takes to set up a microscope optimally and how to tune it for highest performance. One of the «holy grails» of 2

Figure 1.3: (A) Images acquired by transmission electron cryo microscopy of F-actin decorated with myosin Va motor domains with truncated lever arm, in nucleotide-free Rigor state. The defocus value is 1.7 µm and 5 µm for the insert, respectively. (B) The reconstructed density map of the nucleotide-free complex shows the typical helical symmetry of the filamentous actin-backbone. Shown are six actin molecules in their ribbon representation (light yellow) and four myosin motor domains (blue). Images from our own paper [7]. transmission electron microscopy, the Contrast-Transfer-Function (CTF), should then be something, which has lost its legendary aura. While we provide a guideline for experiments with our samples (which we provide as well), we also will give you samples for «free microscopy». This will be tissue samples from biological specimens, plant root or mouse muscle prepared with different protocols. Besides all the proposed experiments, documentation, suggested reading, and discussion, we would like to encourage you to simply «play around» with these samples – following the rule that «one can see a lot just by looking». To make this worthwhile – however – we will discuss your images (and your documentation of this free microscopy) with you and also explain a bit of the biology of the samples.

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2 2.1

Outline of Experiments Day 1

Basic operation and alignment of the transmission electron microscope (TEM) • Get to know the lens system of the microscope. • Align the microscope. • Introduce a sample into the beam path: holey carbon film. Judging image quality • Acquire images of the holey carbon film. • Test the effects of astigmatism and over/under focus on images of holes in the film. • Coherence and Fresnel diffraction.

2.2

Day 2

Diffraction Experiments • Switch the microscope to diffraction mode. • Understand how the patterns are formed and what they tell you about the sample. • Acquire diffraction patterns of thin films of Au, Au/Pd and Pt/Pd. • Use Au patterns to calibrate the camera length for the diffraction patterns. Basic Principles of Bright Field and Dark Field Microscopy • Use the MgO sample to understand how imaging works in the TEM. • Acquire diffraction patterns and bright/dark field images of the MgO crystals.

2.3

Day 3

Bright field Imaging and Effects of the Contrast-Transfer-Function (CTF) • Acquire images of the cross-grating at all (possible) magnification settings to calibrate them. • Measure and use the periodicity of the catalase crystals to calibrate higher magnifications. • Acquire a defocus series of catalase crystals. • Free microscopy.

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2.4

Questionnaire

Be prepared to answer and discuss the following questions over the course of the three days: 1. Why electrons? Discuss the advantages electron microscopy has over light microscopy. 2. Draw the path of the electron beam through the lenses of the microscope. Explain: (a) Condenser, (b) Objective, (c) Projective lens. 3. Explain the following apertures. Where are they located and what is their purpose? (a) Condenser aperture (b) Objective aperture (c) Selected-area aperture 4. What are possible lens aberrations and how can they be compensated? 5. Calculate the relativistic wave length and velocity of electrons for 60 and 80 kV. 6. Why does electron microscopy have to be performed in vacuum? 7. Diffraction / Diffraction Contrast 8. Explain coherence. 9. Bragg/Laue/Ewald - what are these names related to? 10. How is an image formed in the TEM? 11. Bright-field / Dark-field images 12. Point-Spread-Function (PSF) vs. Contrast-Transfer-Function (CTF)

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3

Expected Documentation of the Experiments

Always take note of the illumination conditions at which you record your images (e.g. spotsize, magnification, exposure time, apertures used, ...) to ensure reproducibility and comparability, and minimize errors . When inserting images into your documentation make sure they have a scale bar.

3.1

Day 1 • Explain the lens system of the microscope. • Explain the difference of over and under focus using images of holes in carbon film. • What is astigmatism? Show an example. • Show the effect of beam current, illumination angle and different imaging parameters (e.g. recording times) on the visibility of Fresnel diffraction patterns inside a hole in a carbon film.

3.2

Day 2 • What is diffraction? Consider Bragg and Laue diffraction. • Which lenses are needed for which imaging mode? • Index the diffraction pattern of gold. • Calibrate the camera length. • Use the calibration to analyze the diffraction patterns of Pt/Pd and Au/Pd. • Discuss possible errors. • Show an example of bright-field/dark-field images. Explain what you see.

3.3

Day 3 • Create a calibration table for all magnification settings. How is this done? Discuss possible errors. • Look at Catalase, a protein crystal prepared as negative stain sample, explain «negative stain». • Describe different imaging conditions, show images and their calculated Power Spectrum. What can be seen? What are «Thon rings» (CTF) • Images of «free microscopy», what can be seen?

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4

Experimental Procedure

4.1

Microscope Alignment – The EM109EL

4.1.1

Turning the Microscope on

The microscope can be operated using the control panel and software on the microscope computer. After turning on the microscope electronics (under the table), you can boot up the microscope computer (on the right) and start the TEM control software. Pressing the button POWER in the software will start the microscope and the vacuum pumps. The schematic under the «Vacuum» tab shows you the valves, pumps and sensors. Now you can also boot up the camera computer (left). Once everything is green and the value of P2 is 5 · 10−6 mbar or better (lower), you have clearance to turn on the HIGH TENSION: start with 40 kV , once there, select the next value. Do not be alarmed when the ramp-up exceeds the chosen value. It will go back down. Again, a green bar will indicate when the high tension ramp-up is done. Do this until you have reached 80 kV . Now you have clearance to start the filament by pressing the FILAMENT button in the software. Do not go to high emissions, start with emission setting «2». Closely watch the ramp-up in the software. A drop in vacuum and emission current is indicative of a breakdown. Once the filament has ramped up, you can open the valve on the side of the column (V3), see 4.1. Make sure the light next to it is blinking green. A red flash means you do not yet have clearance to open the valve. To open it, move it to the right and down. The light will now be constantly green. Turn off the light in the room so you can see the fluorescent screen better.

Figure 4.1: The Zeiss EM109EL. Condenser aperture (CA), objective aperture (OA), selected-area aperture (SA), as well as important components are indicated.

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Figure 4.2: Control panel for the EM109EL. 4.1.2

The Control Panel

Figure 4.2 shows the control panel of the EM109EL. The trackball on the left operates the goniometer, shifting the sample position. The sample position can be locked, saved and recalled with the buttons above the trackball. A list of saved positions can be accessed in the GONIOMETER tab of the software. The button F2 will set the filament current control to the trackball. To the right of the trackball is the focus wheel. This will let you focus the image of your sample. The button F3 on the top right of the panel switches between coarse and fine focusing. The magnification of the microscope can be changed with the - and + button above the Focus knob. The current magnification value is shown in Display 1. Here you will also see the objective current while focusing. Pressing Cal will reset the objective and C2 lens. Using the SPOTSIZE knob you can change the illumination, the buttons above can change the emission current. Any changes made here are shown in Display 2. On the far right of the panel are the X/Y knobs. Using the arrow buttons you can choose different settings which can then be controlled using X and Y. The buttons POWER and STANDBY need to be pressed for one second to switch off the microscope or bring it into standby mode (filament and high tension off). It is advised to use the software to control turning everything off. DIFF switches between imaging and diffraction mode of the microscope. 4.1.3

Beam Alignment

Make sure there is no specimen in the beam path when aligning the microscope. Move the slider of the air lock to parking position if it is not there yet (see section 4). Remove the cover of the viewing chamber. The green fluorescence is the electron beam. Set the magnification to 3000x and use the arrow buttons to select BEAM ALIGN to move the beam to the center of the viewing screen. Change the SPOTSIZE and watch the illuminated area: turning the wheel clockwise will decrease the illumination (intensity). Turn it counter-clockwise and find the cross-over (when the brightly illuminated area is smallest). Press FILAMENT (F2) on the panel and use the trackball to lower the emission until you see a «cat’s eye» (turn left! – make sure you do not increase the emission current. Setting it too high will damage the filament. ). If you cannot find 8

Figure 4.3: (Objective) aperture drive. Turning 1 will let you insert or remove the different sized apertures. The black knobs (2 and 3) let you move the aperture in x and y direction. it, try varying the Spotsize again. This can be an iterative process. Set FILAMENT PRECENTERING to the X/Y knobs and try to get an evenly illuminated «cat’s eye». Set BEAM ALIGN to X/Y to center the beam on the viewing screen. The small viewing screen can be inserted using the wheel on the right of the microscope. It has a spot indicating the center. You may have to adjust PRECENTERING again. When done, increase the emission until the beam is saturated (or set it back to the starting value). You may have to press FILAMENT again, since the filament adjustment is turned off after 1 minute. Turn it off, when you are done. Turn the SPOTSIZE knob counter-clockwise past the cross-over. If the beam is astigmatic, you should see a fourfold star. Set C2 STIGMATOR to the X/Y buttons. The astigmatism will be reduced when you see a threefold star (Mercedes star) and the illumination is round. Turn the SPOTSIZE knob until you reach the cross-over. To limit the illuminated area of your sample and decrease lens aberrations, insert the condenser aperture (CA). 4.3 shows an aperture drive. Insert the aperture by turning the silver wheel clockwise. Watch the viewing screen while doing so. You can vary the position of the aperture in two directions by carefully turning the black knobs on the front (2) and side (3). The aperture is centered around the beam when the illuminated area opens concentrically while varying the SPOTSIZE. When you have centered the aperture, turn the SPOTSIZE clockwise to increase the illuminated area. You want to work on the «right side» of the cross-over. 4.1.4

Camera Flat-Field Correction

Before you can use the camera, dark and gain references need to be recorded to compensate for e.g. dirt on the detector. Make sure that there is no specimen in the beam path. Boot up the camera computer and start the software «ImageSP». You will start with recording background (dark) images. For this purpose, you need to make sure that no electrons hit the CCD camera. First, turn on the beam blanker («Pre Specimen») in the TEM software, close valve V3 and put the lid on the viewing chamber to 9

ensure complete darkness. Then insert the camera into the beam path by turning the black lever on the camera (right side of microscope) downwards. Under «Image» you will find a reference image wizard that guides you through the process. The camera software will record dark images for different settings. When it tells you to prepare for gain references, turn the beam blanker off and open the valve. Remove the camera from the beam path to check that you are illuminating the screen evenly. The size of the CCD is about the same as the long edge of the small viewing screen. Make sure the illumination is not too bright before inserting the camera again. Rather start out too dark than too bright. The wizard will start a «Search» window which lets you see the illuminated CCD. In-/decrease the brightness until you have about 6000 counts registered (decrease: clockwise). Electron counts that are much higher than this will damage the CCD. Continue the wizard by clicking «Next». Once you are done creating reference images, check their quality by recording an evenly illuminated image. Choose a short exposure time in the box under the menu bar (200 ms). To the left of the exposure time are three buttons. The rabbit icon opens a «search» window, the video camera gives a live image at the given exposure time value and the camera records a single image. Remove the camera from the beam path before you exchange samples. Make sure not to damage the CCD by recording bright images for long periods of time. NEVER let the direct beam of a diffraction pattern hit the CCD!! Turn the camera out of the beam path when not in use. 4.1.5

Inserting a Sample

TEM sample holders can fit one round sample of 3.05 mm diameter. In transmission electron microscopy, samples need to be thin enough to be electron transparent, i.e. in the order of nanometers in thickness. Samples can be films or dried solutions that have been applied to metal grids, or pieces of bulk samples which have been (embedded and) cut and then thinned down to fit the requirements. To continue the alignment procedure, you will insert a sample of thin carbon film. Practice handling copper grids with the tweezers first, so you do not destroy your sample. Remember that the sample will be inserted into a high vacuum system. Make sure that there is no dirt or lint on the sample and holder which you could introduce into the microscope, thus compromising the vacuum or possibly obstructing the electron beam. Always use gloves when handling the sample holder. 4.4(A) shows the airlock of the microscope and B the necessary tools you need to remove the specimen cartridge (2) from the TEM and exchange the sample. The bottom arrow in (A) points to the specimen slider, here in parking position, meaning the sample is not on the optical axis. 4.4(B) shows the specimen holder after it has been removed and the tools you will need for sample exchange. To remove the cartridge, turn the camera out of the beam path, close the valve V3, turn off the filament and move the slider all the way to the parking position if it is not already there (indicated by the arrow in 4.4 (A)). You will need to press the red button to move the slider. There is one stop halfway between parking position and the optical axis where you need to press the red button again to move the slider. From the parking position, you need to pull and turn the slider to the 12 o’clock position and let it come to rest in the gap there. Now you can turn the airlock counter clockwise (top arrow) until the hole is on the bottom and the specimen cartridge is visible. Screw in the removal tool (1), pull out the cartridge and place it on the holder (3). Make sure you NEVER touch the cartridge! Close the airlock while you exchange the sample. Use the fork tool (5) to lift up the front end of the cartridge and secure it (6). Now 10

Figure 4.4: (A) Air lock. Top Arrow indicates direction to turn when opening the air lock. Bottom arrow indicates the slider, here in parking position. (B) Specimen holder and tools. (C) Specimen Holder when unscrewing the cap. you can unscrew the cap (4) using the tool (7) as shown in 4.4(C). Move the patterned part down to tighten the tool around the cap. Check that there is no grid stuck in the cap. Use the tweezers (8) to carefully remove the specimen grid and exchange it for the carbon film sample. Screw the cap back on and slowly lower the cartridge front using the fork tool. Check that there is no dust on the cartridge, which you could introduce into the microscope, thus compromising the vacuum or blocking the electron beam. Open the airlock again and insert the cartridge. There is a sort of pin on the top of the cartridge when you look at it. This needs to be at around 1 o’clock when inserting the cartridge, you will see a gap to guide you. Push it all the way in, unscrew the tool and use it to check that the cartridge is pushed all the way back. Close the airlock. Press the «Lock in» button to start the evacuation process. This can be done via the software or the AIRLOCK button on the panel. Watch the vacuum system on the software while doing this. There is a list of commands in the left of the microscope schematic. When the software shows a yellow light next to «Turn», turn the slider (red button!) to the 3 o’clock position and wait. If you do not do this quickly enough, the evacuation process will be terminated and you have to start over. Be careful not to cause a vacuum collapse. Clearance is granted once the green light next to «Move» is on and you hear two clicks. Wait until the vacuum has again reached 5 · 10−6 mbar or better, move the sample to the parking position. Restart the filament and watch the ramp up. Open V3 and check the beam alignment (is it astigmatic? - correct C2 STIG if necessary). Move the sample from the parking position onto the optical axis. You should now see your sample. 4.1.6

Adjusting the Objective Astigmatism – Carbon Film Sample

Go to a low magnification mode to get an overview of your sample. Use the trackball on the panel or choose GONIO on the X/Y knobs to move the sample around. You 11

should see the bars of a copper (or other metal) grid. They will be very dark, because they are too thick to be electron transparent. The (very thin) carbon film will be on some sort of plastic support on the copper grid. This can be a lacy carbon film, which has holes of different sizes, or a Quantifoil, which has equidistant holes of one size. If the carbon film is intact, covering the whole grid, you will not be able to see it. Look for holes or tears in the carbon to find areas covered in carbon. Choose a large, carbon-covered hole in your lacy film. Move the edge of your film into the middle of the screen. Increase the magnification and bring the sample into focus. What happens when you focus/under focus/over focus the image? Find an area of (only) carbon film. If there are some dirt particles on it, use them to bring the sample into focus. Go to a magnification of 85000x and make sure the screen is evenly illuminated, but not too bright. Insert the camera and start the “Search” mode (rabbit button). Make sure you do not have more than 6000 counts on the camera. This is displayed below the histogram on the left in the camera software. If you see spots in the FFT you are overexposing the camera. This can damage the CCD. Decrease the illumination by turning the Spotsize knob clockwise or decrease the exposure time of the camera in the menu bar. You should be able to see some sort of ring pattern in the diffractogram (FFT) when under/over focusing your image. Discuss what this means. How can you differentiate between under- and over focus using the FFT of your image? Use the ring pattern in under focus to adjust the Objective Stigmator (OBJ STIG). You have corrected the astigmatism when the rings are perfectly round. If you cannot see a ring pattern, use the image of a round hole in your sample to adjust the objective astigmatism. How can you tell when you have corrected the astigmatism, just from the image of the hole? Use the holes of the lacy carbon to investigate Fresnel diffraction. Vary recording times and illumination conditions for this.

4.2

Electron Diffraction

Insert the sample grid with the gold film into the microscope. Choose a sample area and bring the sample into focus as well as you can. The crystals are very small. Use a hole in the film to adjust focus. Go to the highest magnifications if you want to see the crystals but it is hard to get them into focus. To acquire diffraction images, you do not need to be at highest magnification. At a magnification of 20000x you can see the smallest selected-area aperture (SA) well. Make sure you have removed the objective aperture. Switch to diffraction mode with the DIFF button on the panel. Vary the illumination until the center spot is focused to a sharp point. Explain what you see and why you see it. Insert the selected-area aperture. What happens? What can a diffraction pattern tell you about the sample you are investigating? Go to imaging mode to center the SA aperture. Bring the diffraction pattern into focus with the SPOTSIZE knob. Since the camera is situated on the side of the microscope, we do not have the option of blocking the direct beam in the diffraction pattern. Its high intensity will damage the CCD chip of the camera, thus, we cannot record a full diffraction pattern. Insert the largest objective aperture again. Move it to one side, so that you just do not see the direct beam anymore. Record an image. Repeat this on the other side of the diffraction pattern. Analysis is easier if you record images like this around the whole diffraction pattern. Make sure you do not let the direct beam hit the camera! If your diffraction pattern is too bright for the CCD, choose a smaller aperture. Average the images into one using ImageJ/Fiji and calculate the real camera lengths using the 12

known values for gold. Compare them to the nominal camera lengths. The diffraction pattern of gold will fit well on the CCD for camera lengths of 150 and 250 mm. 400 mm or more might not fit anymore and, thus, cannot be calibrated. Make sure you always use the same imaging conditions and make a note of them in your lab journal. Once you have acquired your calibration measurements, exchange the sample for the grid covered in Pt/Pd (Au/Pd). Record the diffraction pattern in the same manner using the objective aperture. Use Fiji/ImageJ to merge the images and measure the lattice spacings of the crystals.

4.3

Magnesium Oxide

Insert the magnesium oxide sample. Get an overview of what you are looking at. What can you see? Choose an area with several MgO crystals and increase the magnification. Use the objective aperture to record bright field and dark field images. Explain what you see and why you see it. What happens when you acquire images in areas of the diffraction pattern that are across from each other? Why? Record dark field images for several different aperture positions at different magnifications.

4.4

Calibration

Calibration measurements need to be done before you can start analyzing images of your samples in the microscope. For this purpose a special calibration standard will be used: a cross-grating (2160 lines/mm) with latex spheres (Ø 0.261 µm). Go to the lowest magnification and bring the sample into focus. What do you see? Switch to higher magnifications, adjust the spotsize and focus while doing so. What happens to the image? Why does it rotate? Switch to diffraction mode. Center the largest objective aperture around the direct beam and switch to imaging mode. What happens to the image? Calibrate each magnification using the cross-grating and check your values using the latex spheres. Create a calibration table and discuss the difference between nominal and real magnification. Make sure your sample is always in focus when calibrating a magnification. Pressing the «nm» button on the bottom of the image window in the camera opens a calibration tool. You can save calibrations in the software but it is more reliable to use Fiji/ImageJ to create your own calibration table in your lab journal. As long as you see the repeating pattern of the cross-grating, you can use the FFT of the image to do the calibration, otherwise use a line scan profile. You will not be able to calibrate all magnifications due to the limited size of the periodical cross-grating pattern.

4.5

Catalase

Exchange the grid for a sample containing bovine catalase[8]. The catalase you will see in the TEM will be in crystalline form. Find the catalase crystals, of rectangular form, on the sample. Increase the magnification. Acquire images of the catalase crystals. When going to higher magnification you should be able to see the lattice spacings of the catalase crystal in the image as well as in the FFT. Measure the spacings of the catalase and use the values to calibrate the higher magnifications that could not fit the cross-grating lines. Add the new magnification values to your table. Use the catalase to investigate different imaging conditions ((over/under) focus, filament heating, ...) and the CTF. 13

5

Helpful Features of Fiji

Fiji (Fiji is just ImageJ) is a scientific image processing software[9]. You can use it to calibrate and analyze electron microscopy images. We will list some features to help you with your experiment.

5.1

Opening images

You can open several images as one stack under File – Import – Image Sequence. Fiji cannot read any calibration done by this camera software. Reset the image size to pixels for each image under Image – Properties (pixel: 1x1) when doing calibrations or the calibrated pixel size in nm, once you know it.

5.2

Fast Fourier Transforms

Under Process – FFT – FFT you can let Fiji calculate the FFT of your image or of a selected part of your image. This can help you calibrate your magnifications.

5.3

Image Stacks

Image stacks can be created from sequential images. Under Image – Stacks – z-Project you can create a projection of the whole image stack. When putting together the recorded partial diffraction patterns, choosing «Max Intensity» will let you create a single image containing the brightest parts of each single image.

5.4

Scale Bars

Make sure all your images have scale bars. You can insert a scale bar after you have set your pixel size under Analyze - Tools - Scale Bar. Make sure it is visible.

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Provided Samples • Holey carbon film • Gold (Au) • Gold/Palladium (Au/Pd 80:20) • Platinum/Palladium (Pt/Pd 80:20) • Magnesium oxide crystals • Cross-grating calibration sample with latex spheres (2160 lines/mm, ø 0.261µm) • Bovine catalase • Samples for «free microscopy», will vary from group to group, usually samples of plant root material and/or tissue samples, e.g. from mouse muscle.

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References / Suggested Reading [1] David B. Williams and C. Barry Carter. Transmission Electron Microscopy: A Textbook for Materials Science. Springer Science & Business Media, July 2009. ISBN 978-0-387-76501-3. [2] Ludwig Reimer and Helmut Kohl. Transmission Electron Microscopy, volume 36 of Springer Series in Optical Sciences. Springer New York, New York, NY, 2008. ISBN 978-0-387-40093-8 978-0-387-34758-5. URL http://link. springer.com/10.1007/978-0-387-40093-8. [3] Rolf Erni. Aberration-corrected imaging in transmission electron microscopy: an introduction. ICP/Imperial College Press ; Distributed by World Scientific Pub. Co, London : Singapore ; Hackensack, NJ, 2010. ISBN 978-1-84816-536-6. [4] John Rodenburg. The beginner’s guide to transmission electron microscopy. URL http://www.rodenburg.org/guide/index.html. [5] Max Haider, Harald Rose, Stephan Uhlemann, Eugen Schwan, Bernd Kabius, and Knut Urban. A spherical-aberration-corrected 200 kV transmission electron microscope. Ultramicroscopy, 75(1):53–60, October 1998. ISSN 0304-3991. doi: 10. 1016/S0304-3991(98)00048-5. URL http://www.sciencedirect.com/ science/article/pii/S0304399198000485. [6] SALVE Sub Angstrom Low Voltage Electron Microscopy Project III of Ulm University / FEI company / CEOS / DFG / MWK: Startpage. URL http: //www.salve-project.de/. [7] Sarah F. Wulf, Virginie Ropars, Setsuko Fujita-Becker, Marco Oster, Goetz Hofhaus, Leonardo G. Trabuco, Olena Pylypenko, H. Lee Sweeney, Anne M. Houdusse, and Rasmus R. Schröder. Force-producing ADP state of myosin bound to actin. Proceedings of the National Academy of Sciences, 113(13):E1844–E1852, March 2016. ISSN 0027-8424, 1091-6490. doi: 10.1073/pnas.1516598113. URL http://www.pnas.org/content/113/13/E1844. [8] Nicholas G. Wrigley. The lattice spacing of crystalline catalase as an internal standard of length in electron microscopy. Journal of Ultrastructure Research, 24(5-6):454–464, September 1968. ISSN 0022-5320. doi: 10. 1016/S0022-5320(68)80048-6. URL http://www.sciencedirect.com/ science/article/pii/S0022532068800486. [9] Johannes Schindelin, Ignacio Arganda-Carreras, Erwin Frise, Verena Kaynig, Mark Longair, Tobias Pietzsch, Stephan Preibisch, Curtis Rueden, Stephan Saalfeld, Benjamin Schmid, Jean-Yves Tinevez, Daniel James White, Volker Hartenstein, Kevin Eliceiri, Pavel Tomancak, and Albert Cardona. Fiji: an opensource platform for biological-image analysis. Nature Methods, 9(7):676–682, July 2012. ISSN 1548-7091. doi: 10.1038/nmeth.2019. URL http://www. nature.com/nmeth/journal/v9/n7/full/nmeth.2019.html.

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Appendix Diffraction: Gold h 0 1 0 0 1 2 0 1 0 2 3 1 0 1 2

k 0 1 0 2 1 2 0 3 2 2 3 1 4 3 4

l 0 1 2 2 3 2 4 3 4 4 3 5 4 5 4

d* [nm−1 ] 0 4.256 4.914 6.949 8.149 8.511 9.828 10.71 10.988 12.037 12.767 12.767 13.899 14.536 14.742

d [nm] 0.235 0.204 0.144 0.123 0.117 0.102 0.093 0.091 0.083 0.078 0.078 0.072 0.069 0.068

Table 6.1: Peak positions for gold (Au).

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