Name: mitochondria and endoplasmic reticulum imaging by correlative light and volume electron microscopy
Uploaded: 2019-07-20T12:30:00
Description: Watch this Scientific Journal Video about Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy at JoVE.com

This research was supported by KBRI basic research program through Korea Brain Research Institute funded by Ministry of Science and ICT (19-BR-01-08), and National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT)(No. 2019R1A2C1010634). SCO1-APEX2 and HRP-KDEL plasmids were kindly provided by Hyun-Woo Rhee (Seoul National University). TEM data were acquired at Brain Research Core Facilities in KBRI.

1. Cell culture with patterned grid culture dish and cell transfection with SCO1-APEX2 and HRP-KDEL plasmid vector
<ol>
	<li>Seed 1 x 105 HEK293T cells by placing them into 35-mm glass grid-bottomed culture dishes in a humidified atmosphere containing 5% CO2 at 37 &#176;C in Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) supplemented with 10% fetal bovine serum, 100 U/mL penicillin, and 100 U/mL streptomycin.</li>
	<li>The day after seeding the cells, when they have grown to 50%-60% conflue...

<ol>
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Determining the cellular localization of specific proteins at a nanometer resolution using EM is crucial to understand the cellular functions of proteins. Generally, there are two techniques to study the localization of a target protein via EM. One is the immunogold technique, which has been used in EM since 1960, and the other is a technique using recently developed genetically encoded tags16. Traditional immunogold techniques have employed antibody-conjugated gold particles or quantum dots to sh...

Mitochondria and endoplasmic reticulum (ER) are membrane-bound cellular organelles. Their connection is necessary for their function, and proteins related to their network have been described1. The distance between the mitochondria and ER has been reported as approximately 100 nm using light microscopy2; however, recent super-resolution microscopy3 and electron microscopy (EM)4 studies have revealed it to be considerably smaller, at approximately 10-25 nm. The resolution achieved in super-resolution microscopy is lower than EM, and specific labeling is necessary. EM is a suitable technique to attain a sufficiently high-resolution contrast for structural studies of the connections between mitochondria and ER. However, a disadvantage is the limited z-axis information because the thin sections must be approximately 60 nm or thinner for conventional transmission electron microscopy (TEM). For sufficient EM z-axis imaging, three-dimensional electron microscopy (3DEM) can be used5. However, this involves the preparation of hundreds of thin serial sections of whole cells, which is very tricky work that only a few skilled technologists have mastered. These thin sections are collected on fragile formvar film-coated one-hole TEM grids. If the film breaks on one gird, serial imaging and volume reconstruction is not possible. Serial block-face scanning electron microscopy (SBEM) is a popular technique for 3DEM that uses destructive en bloc sectioning inside the scanning electron microscope (SEM) vacuum chamber with either a diamond knife (Dik-SBEM) or a focused ion beam (FIB-SEM)6. However, because those techniques are not available at all facilities, we suggest array tomography7 using serial sectioning and SEM. In array tomography, serial sections cut using an ultramicrotome are transferred to a glass coverslip instead of a TEM grid and visualized via light microscopy and SEM8. To enhance the signal for backscatter electron (BSE) imaging, we utilized an en bloc EM staining protocol employing osmium tetroxide (OsO4)-fixed cells with osmiophilic thiocarbohydrazide (TCH)9, enabling us to obtain images without post-embedding double staining.
Additionally, the mitochondrial marker SCO1 (cytochrome c oxidase assembly protein 1)&#8211; ascorbate peroxidase 2 (APEX2)10 molecular tag was used to visualize mitochondria at the EM level. APEX2 is approximately 28 kDa and is derived from soybean ascorbate peroxidase11. It was developed to show the detailed location of specific proteins at the EM level in the same way that green fluorescent protein-tagged protein is used in light microscopy. APEX2 converts 3,3&#39;&#160;-diaminobenzidine (DAB) into an insoluble osmiophilic polymer at the site of the tag in the presence of the cofactor hydrogen peroxide (H2O2). APEX2 can be used as an alternative to traditional antibody labeling in EM, with a protein localization throughout the depth of the entire cell. In other words, the APEX2-tagged protein can be visualized by specific osmication11 without immunogold labeling and permeabilization after ultra-cryosectioning. Horseradish peroxidase (HRP) is also a sensitive tag that catalyzes the H2O2-dependent polymerization of DAB into a localized precipitate, providing EM contrast after treatment with OsO4. The ER target peptide sequence HRP-KDEL (lys-asp-glu-leu)12 was applied to visualize ER within a whole cell. To evaluate our protocol of utilizing genetic tags and en bloc staining with reduced osmium and TCH (rOTO method), using the osmication effect at the same time, we compared the membrane contrast with and without the use of each genetic tag in rOTO en bloc staining. Although 3DEM with array tomography and DAB staining with APEX and HRP have, respectively, been utilized for other purposes, our protocol is unique because we have combined array tomography for 3DEM and DAB staining for mitochondria and ER labeling. Specifically, we showed five cells with and without APEX-tagged genes in the same section, which aided in investigating the effect of the genetic modification on cells.

<table border="0" cellpadding="0" cellspacing="0" style="width:869px;" width="868">
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	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Glutaraldehyde</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">16200</td>
			<td style="width:349px;">Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Paraformaldehyde</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">19210</td>
			<td>Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Sodium cacodylate</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">12300</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Osmium tetroxide 4 % aqueous solution</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">16320</td>
			<td>Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Epon 812</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">14120</td>
			<td>EMbed 812- 20 ml/ DDSA- 16 ml/ NMA- 8 ml/ DMP-30 - 0.8 ml</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Ultra-microtome Leica ARTOS 3D</td>
			<td style="width:123px;">Leica</td>
			<td style="width:123px;">ARTOS 3D</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Uranyl acetate</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">22400</td>
			<td>Hazardous chemical</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Lead citrate</td>
			<td style="width:123px;">EMS</td>
			<td style="width:123px;">17900</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:275px;">35mm Gridded coverslip dish</td>
			<td style="width:123px;">Mattek</td>
			<td style="width:123px;">P35G-1.5-14-CGRD</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Glow discharger</td>
			<td style="width:123px;">Pelco</td>
			<td style="width:123px;">easiGlow</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Formvar carbon coated Copper Grid</td>
			<td style="width:123px;">Ted Pella</td>
			<td style="width:123px;">01805-F</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Hydrochloric acid</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">258148</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fugene HD</td>
			<td style="width:123px;">Promega</td>
			<td style="width:123px;">E2311</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Glycine</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">G8898</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:275px;">3,3&#8242; -diaminobenzamidine (DAB)</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">D8001</td>
			<td>Hazardous chemical</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">30% Hydrogen peroxide solution</td>
			<td style="width:123px;">Merck</td>
			<td style="width:123px;">107210</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Potassium hexacyanoferrate(II) trihydrate</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">P3289</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">0.22 um syringe filter</td>
			<td style="width:123px;">Sartorius</td>
			<td style="width:123px;">16534</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Thiocarbonyldihydrazide</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">223220</td>
			<td>Use only in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Potassium hydroxide</td>
			<td style="width:123px;">Fluka</td>
			<td style="width:123px;">10193426</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">L-aspartic acid</td>
			<td style="width:123px;">SIGMA</td>
			<td style="width:123px;">A9256</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Ethanol</td>
			<td style="width:123px;">Merck</td>
			<td style="width:123px;">100983</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Transmission electron microscopy</td>
			<td style="width:123px;">FEI</td>
			<td style="width:123px;">Tecnai G2</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:275px;">Indium tin oxide (ITO) coated glass coverslips</td>
			<td style="width:123px;">SPI</td>
			<td style="width:123px;">06489-AB</td>
			<td>fragil glass</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Isopropanol</td>
			<td style="width:123px;">Fisher Bioreagents</td>
			<td style="width:123px;">BP2618-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Diamond knife</td>
			<td style="width:123px;">Leica</td>
			<td style="width:123px;">AT-4</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Inveted light microscopy</td>
			<td style="width:123px;">Nikon</td>
			<td style="width:123px;">ECLipse TS100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Scanning electron microscopy</td>
			<td style="width:123px;">Zeiss</td>
			<td style="width:123px;">Auriga</td>
			<td></td>
		</tr>
	</tbody>
</table>

mitochondria and endoplasmic reticulum imaging by correlative light and volume electron microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

Figure 1 describes the schedule and workflow for this protocol. The protocol requires 7 days; however, depending on the time spent on SEM imaging, this may increase. For cell transfection, the confluency of the cells should be controlled so as not to cover the bottom of the entire grid plate (Figure 1A). A high cell density could prevent the identification of the cell of interest during light microscope and EM observation. We used genetically tagged plasmids that expressed APEX2 and HRP ...

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Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

This volume electron microscopy technique is used to perform biological studies that require the observation of a 3-D structure. Offering a rod, fielder view, and sample storage before image retakes. Fx2 Asan engineered the path stages that catalyzes the deabilation to increase the electron test of target structures and can be used to locate a specific target of interest.These particles combines the use of correlative light and 3-D electron microscopy technique to investigate the interactions between membranous organelles and host cells. 16 to 24 hours after transfection, gently wash the culture with 250 microliters of 30 to 37 degree Celsius fixation solution. Quickly replacing the wash with 1.5 milliliters of fresh fixation solution, and placing the culture on ice for 30 minutes.At the end of the incubation, rinse the cells with three ten minute washes in one milliliter of ice-cold 0.1 molar sodium cacodylate buffer per wash. After the last wash, add one milliliter of ice-cold 0 to 4 degrees Celsius 20 millimolar glycine solution to the dish for a ten-minute incubation on ice followed by three five minute washes with one milliliter of fresh, cold 0.1 molar sodium cacodylate buffer per wash. Next, label the cells with 500 microliters of freshly prepared DAB solution.And incubate the cells on ice for approximately five to forty-five minutes. When a light brown stain is visible under an inverted light microscope, rinse the cells three times with one milliliter of fresh, cold 0.1 molar sodium cacodylate buffer for ten minutes per wash. Then, use a phase contrast inverted microscope to visualize the DAB staining at a 100 times magnification or higher and mark the bottom of the glass where the region of interest is located.For electron microscope block sample preparation, first post-fix the cells with one milliliter of 2%reduced osmium tetroxide for one hour at four degrees Celsius. At the end of the fixation, rinse the cells with three five-minute washes and one milliliter of fresh distilled water per wash. Before treating the cells with one milliliter of filtered TCH solution for 20 minutes.At the end of the incubation, wash the cells three times in distilled water as just demonstrated and fix the cells with a 30-minute incubation in 2%reduced osmium tetroxide. At the end of the second fixation, wash the cells three times with distilled water and cover the culture with one milliliter of aqueous 1%uranyl acetate overnight at 4 degrees Celsius protected from light. The next morning, wash the cells three times with distilled water before staining with one milliliter of 60 degree Celsius warmed Walton's lead aspartate solution for 30 minutes in a 60 degrees Celsius oven.At the end of the incubation, wash the cells three times in distilled water. And incubate the culture in a graded series of 20-minute 2 milliliter ethanol aliquots. After the last 100%ethanol exposure, incubate the cells for 30 minutes in one milliliter of three to one ethanol to low viscosity embedding mixture medium at room temperature.At the end of the incubation, replace the medium with one milliliter of one-to-one ethanol to low viscosity embedding mixture medium for another 30 minute incubation at room temperature. Next, replace the medium with one millilter of one-to-three ethanol to low viscosity embedding mixture medium for an additional 30 minutes at room temperature. Then, replace the medium with one milliliter of 100%low viscosity embedding medium overnight, followed by embedding of the sample in 100%low viscosity embedding mixture for 24 hours at 60 degrees Celsius.For transmission electron microscopy analysis, the next day, use an ultra microtome to obtain 90 nanometer thick sections and observe the grid under transmission electron microscopy at 200 kilovolts. Before beginning the sample preparation, clean Indium Tin Oxide coated glass coverslips with gentle agitation in isopropanol for 30 to 60 seconds. At the end of the agitation, drain off the excess alcohol and place the coverslips in a dust-free environment.When the coverslips are dry, use a plasma coater to glow discharge the coverslips for one minute. Then insert the coverslips into the substrate holder and place the substrate holder into the knife boat. To obtain samples for scanning electron microscopy, insert the embedded sample block into the sample holder of the ultramicrotome.And set the holder into the trimming block. Use a razor blade to trim away the excess resin around the target position so that the face of the block acquires a trapezoid or rectangular shape. The leading and trailing edges should be strictly parallel to each other.It is not easy to acquire a large number of serial sections. However, you have to have the leading edges and trailing edges completely parallel. Next, insert the sample holder into the arm of the ultramicrotome.Place the diamond knife in the knife holder, and fill the knife boat with filtered distilled water. Then, carefully push the handle of the holder to adjust the carrier position so that the edge of the coverslips is close to the knife. After obtaining the sample, stop the sectioning process, slowly open the clamping screw of the tube and drain the water boat.When the ribbon collection process is complete, remove the substrate with the handle of the clamping device and dry the ribbon. The use of genetically tagged plasmids expressing proteins of interest, allows the identification of the transfected target cells after staining for the tagged proteins. Correlative light electron microscopy can then be used to further visualize the labeled cell culture.For example, the dark stain generated by SCO1-APEX2 is observed exclusively in the mitochondrial intermembrane space and not in the matrix space of the mitochondria. Code transfected cells with both SCO1-APEX2 and HRP-KDEL plasmids express a highly dense electron signal in the mitochondrial intermembrane space and the endoplasmic reticulum. However, no endoplasmic reticulum staining is observed in cells that are transfected with SCO1-APEX2 only.For serial imaging by scanning electron microscopy, an overview of the whole array image is created using a backscatter electron detector over a large area. Next, the region of interest is placed in the first section and propagated to all the other sections. Finally, the region of interest containing five target cells is imaged with five nanometer imaged pixels.Magnification reveals detailed subcellular structures such as the Golgi apparatus, mitochondria, and endpolasmic reticulum. Serial imaging clearly reveals that endoplasmic reticulum-mitochondria contacts occur on different z-planes. You should have a good understanding of how to locate a specific target structure using correlative light and electron microscopy to investigate the effects of a specific identity modification.These staining techniques combined with volume electron microscopy could be used for larger scale EM to investigate cellular interactions particularly in eurovirus. Remember the glutaraldehyde, also enteterocide and TCH are very toxic, so be sure to wear protective clothing and to work in a fume hood when using these materials.

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Unit 1

Transmembrane Transport in Endoplasmic Reticulum and Peroxisomes

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Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Imaging Amyloid Tissues Stained with Luminescent Conjugated Oligothiophenes by Hyperspectral Confocal Microscopy and Fluorescence Lifetime Imaging

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease afflicting primarily the central nervous system, and systemic amyloidosis where serum amyloid A, transthyretin and IgG light chains deposit as amyloid in liver, carpal tunnel, spleen, kidney, heart, and other peripheral tissues. Amyloid has been known and studied for more than a century, often using amyloid specific dyes such as Congo red and Thioflavin T (ThT) or Thioflavin (ThS). In this paper, we present heptamer-formyl thiophene acetic acid (hFTAA) as an example of recently developed complements to these dyes called luminescent conjugated oligothiophenes (LCOs). hFTAA is easy to use and is compatible with co-staining in immunofluorescence or with other cellular markers. Extensive research has proven that hFTAA detects a wider range of disease associated protein aggregates than conventional amyloid dyes. In addition, hFTAA can also be applied for optical assignment of distinct aggregated morphotypes to allow studies of amyloid fibril polymorphism. While the imaging methodology applied is optional, we here demonstrate hyperspectral imaging (HIS), laser scanning confocal microscopy and fluorescence lifetime imaging (FLIM). These examples show some of the imaging techniques where LCOs can be used as tools to gain more detailed knowledge of the formation and structural properties of amyloids. An important limitation to the technique is, as for all conventional optical microscopy techniques, the requirement for microscopic size of aggregates to allow detection. Furthermore, the aggregate should comprise a repetitive &#946;-sheet structure to allow for hFTAA binding. Excessive fixation and/or epitope exposure that modify the aggregate structure or conformation can render poor hFTAA binding and hence pose limitations to accurate imaging.

Amyloid deposition is a hallmark of different diseases and afflicts many different organs. This paper describes the application of luminescent conjugated oligothiophene fluorescence staining in combination with fluorescence microscopy techniques. This staining method represents a powerful tool for detection and exploration of protein aggregates in both clinical and scientific setups.

Proteins that deposit as amyloid in tissues throughout the body can be the cause or consequence of a large number of diseases. Among these we find ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Analysis of &amp;#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

Imaging of Extracellular Vesicles by Atomic Force Microscopy

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane proteins and other molecules, and diverse luminal content inherited from a parent cell, which includes RNAs, proteins, and DNAs. The characterization of the hydrodynamic sizes of EVs, which depends on the size of the vesicle and its coronal layer formed by surface decorations, has become routine. For exosomes, the smallest of EVs, the relative difference between the hydrodynamic and vesicles sizes is significant. The characterization of vesicles sizes by the cryogenic transmission electron microscopy (cryo-TEM) imaging, a gold standard technique, remains a challenge due to the cost of the instrument, the expertise required to perform the sample preparation, imaging and data analysis, and a small number of particles often observed in images. A widely available and accessible alternative is the atomic force microscopy (AFM), which can produce versatile data on three-dimensional geometry, size, and other biophysical properties of extracellular vesicles. The developed protocol guides the users in utilizing this analytical tool and outlines the workflow for the analysis of EVs by the AFM, which includes the sample preparation for imaging EVs in hydrated or desiccated form, the electrostatic immobilization of vesicles on a substrate, data acquisition, its analysis, and interpretation. The representative results demonstrate that the fixation of EVs on the modified mica surface is predictable, customizable, and allows the user to obtain sizing results for a large number of vesicles. The vesicle sizing based on the AFM data was found to be consistent with the cryo-TEM imaging.&#160;

A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties.&#160;

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...

Single Particle Cryo-Electron Microscopy: From Sample to Structure

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The overall process involves i) vitrifying the specimen in a thin film supported on a cryoEM grid; ii) screening the specimen to assess particle distribution and ice quality; iii) if the grid is suitable, collecting a single particle dataset for analysis; and iv) image processing to yield an EM density map. In this protocol, an overview for each of these steps is provided, with a focus on the variables which a user can modify during the workflow and the troubleshooting of common issues. With remote microscope operation becoming standard in many facilities, variations on imaging protocols to assist users in efficient operation and imaging when physical access to the microscope is limited will be described.

Structure determination of macromolecular complexes using cryoEM has become routine for certain classes of proteins and complexes. Here, this pipeline is summarized (sample preparation, screening, data acquisition and processing) and readers are directed towards further detailed resources and variables that may be altered in the case of more challenging specimens.

Cryo-electron microscopy (cryoEM) is a powerful technique for structure determination of macromolecular complexes, via single particle analysis (SPA). The ...

Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules present in the cellular context and preserved in a near-native frozen-hydrated state. However, there are challenges associated with culturing and/or adhering cells onto TEM grids in a manner that is suitable for tomography while retaining the cells in their physiological state. Here, a detailed step-by-step protocol is presented on the use of micropatterning to direct and promote eukaryotic cell growth on TEM grids. During micropatterning, cell growth is directed by depositing extra-cellular matrix (ECM) proteins within specified patterns and positions on the foil of the TEM grid while the other areas remain coated with an anti-fouling layer. Flexibility in the choice of surface coating and pattern design makes micropatterning broadly applicable for a wide range of cell types. Micropatterning is useful for studies of structures within individual cells as well as more complex experimental systems such as host-pathogen interactions or differentiated multi-cellular communities. Micropatterning may also be integrated into many downstream whole-cell cryo-ET workflows, including correlative light and electron microscopy (cryo-CLEM) and focused-ion beam milling (cryo-FIB).

The goal of this protocol is to direct cell adhesion and growth to targeted areas of grids for cryo-electron microscopy. This is achieved by applying an anti-fouling layer that is ablated in user-specified patterns followed by deposition of extra-cellular matrix proteins in the patterned areas prior to cell seeding.

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules ...

Extracting Modified Microtubules from Mammalian Cells to Study Microtubule-Protein Complexes by Cryo-Electron Microscopy

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the posttranslational modifications, microtubules can form complexes with various interacting proteins. These microtubule-protein complexes are often implicated in human diseases. Understanding the structure of such complexes is useful for elucidating their mechanisms of action and can be studied by cryo-electron microscopy (cryo-EM). To obtain such complexes for structural studies, it is important to extract microtubules containing or lacking specific posttranslational modifications. Here, we describe a simplified protocol to extract endogenous tubulin from genetically modified mammalian cells, involving microtubule polymerization, followed by sedimentation using ultracentrifugation. The extracted tubulin can then be used to prepare cryo-electron microscope grids with microtubules that are bound to a purified microtubule-binding protein of interest. As an example, we demonstrate the extraction of fully tyrosinated microtubules from cell lines engineered to lack the three known tubulin-detyrosinating enzymes. These microtubules are then used to make a protein complex with enzymatically inactive microtubule-associated tubulin detyrosinase on cryo-EM grids.

Here, we describe a protocol to extract endogenous tubulin from mammalian cells, which can lack or contain specific microtubule-modifying enzymes, to obtain microtubules enriched for a specific modification. We then describe how the extracted microtubules can be decorated with purified microtubule-binding proteins to prepare grids for cryo-electron microscopy.

Microtubules are an important part of the cytoskeleton and are involved in intracellular organization, cell division, and migration. Depending on the ...

Accelerating Macromolecular Structural Analysis Using Smart Leginon Autoscreen

Cryo-Electron Microscopy Screening Automation Across Multiple Grids Using Smart Leginon

Advancements in cryo-electron microscopy (cryoEM) techniques over the past decade have allowed structural biologists to routinely resolve macromolecular protein complexes to near-atomic resolution. The general workflow of the entire cryoEM pipeline involves iterating between sample preparation, cryoEM grid preparation, and sample/grid screening before moving on to high-resolution data collection. Iterating between sample/grid preparation and screening is typically a major bottleneck for researchers, as every iterative experiment must optimize for sample concentration, buffer conditions, grid material, grid hole size, ice thickness, and protein particle behavior in the ice, amongst other variables. Furthermore, once these variables are satisfactorily determined, grids prepared under identical conditions vary widely in whether they are ready for data collection, so additional screening sessions prior to selecting optimal grids for high-resolution data collection are recommended. This sample/grid preparation and screening process often consumes several dozen grids and days of operator time at the microscope. Furthermore, the screening process is limited to operator/microscope availability and microscope accessibility. Here, we demonstrate how to use Leginon and Smart Leginon Autoscreen to automate the majority of cryoEM grid screening. Autoscreen combines machine learning, computer vision algorithms, and microscope-handling algorithms to remove the need for constant manual operator input. Autoscreen can autonomously load and image grids with multi-scale imaging using an automated specimen-exchange cassette system, resulting in unattended grid screening for an entire cassette. As a result, operator time for screening 12 grids may be reduced to ~10 min with Autoscreen compared to ~6 h using previous methods which are hampered by their inability to account for high variability between grids. This protocol first introduces basic Leginon setup and functionality, then demonstrates Autoscreen functionality step-by-step from the creation of a template session to the end of a 12-grid automated screening session.

Author Spotlight: Enhancing Cryo-Electron Microscopy by Automated Data Collection and Analysis Techniques

Cryo-electron microscopy (cryoEM) multi-grid screening is often a tedious process that demands hours of attention. This protocol shows how to set up a standard Leginon collection and Smart Leginon Autoscreen to automate this process. This protocol can be applied to the majority of cryoEM holey foil grids.

Advancements in cryo-electron microscopy (cryoEM) techniques over the past decade have allowed structural biologists to routinely resolve macromolecular ...