The authors wish to thank Fabienne Proamer, Jean-Yves Rinckel, David Hoffmann, Monique Freund for technical assistance. This work was supported by ARMESA (Association de Recherche et D&#233;veloppement en M&#233;decine et Sant&#233; Publique), the European Union through the European Regional Development Fund (ERDF) and by Grant ANR-17-CE14-0001-01 to H.d.S.

All animal experiments were performed in accordance with European standards 2010/63/EU and the CREMEAS Committee on the Ethics of Animal Experiments of the University of Strasbourg (Comit&#233; R&#233;gional d&#39;Ethique en Mati&#232;re d&#39;Exp&#233;rimentation Animale Strasbourg). The protocol is schematically shown in Figure 1.
1. Bone marrow collection and fixation (&#160;Figure 1A)
CAUTION: This procedure includes carcinogenic, mutagenic, and/or toxic substances and is performed under a chemical extraction hood. Wear appropriate protective equipment such as gloves and protections glasses.
<ol>
	<li>Prepare the fixative solution consisting of 2.5% glutaraldehyde in cacodylate buffer (see Supplementary File).</li>
	<li>Bone marrow collection
	<ol>
		<li>Use adult C57BL/6 mice of either sex 12-18 weeks of age. Euthanize the mice by CO2 asphyxiation and cervical dislocation.</li>
		<li>With a pair of thin scissors, cut the skin around the thigh and use tweezers to peel the skin off. Remove the extremity of the paw and then cut between the hip and thigh. Detach tibia from femur by cutting at the knee articulation and remove adherent tissue on tibias and femurs by using a scalpel.</li>
		<li>Remove the epiphyses with a sharp razor blade. While holding the femur with tweezers, use a 5 mL syringe filled with cacodylate buffer with a 21 G needle to flush the bone marrow into a 15 mL tube filled with 2 mL cacodylate buffer. To do so, insert the bevel of the needle into the bone marrow opening and slowly press the plunger until the marrow is expelled.</li>
	</ol>
	</li>
	<li>Bone marrow fixation by rapid immersion into fixative.
	<ol>
		<li>Immediately after flushing, use a plastic pipette to transfer the bone marrow cylinder into 1 mL of fresh glutaraldehyde fixative solution (previously prepared in 1.1) for 60 min at room temperature. 
		&#8203;NOTE: To preserve the tissue, ensure that the entire process, from bone dissection to the fixation step, is completed in less than 10 min. For the fixation, ensure that the fixative solution is at room temperature to avoid heat shock.</li>
	</ol>
	</li>
</ol>
2. Embedding bone marrow in agarose
NOTE: Marrow tissue is not sufficiently cohesive to maintain its integrity during the different washing steps and material can be easily lost. To overcome this problem, the marrow is covered in a gel of agar before dehydration.
<ol>
	<li>Prepare the agarose solution as described in the Supplementary File.</li>
	<li>Wash the fixed marrow from section 1.3 in cacodylate buffer and transfer it carefully to a glass slide using a plastic pipette. Using a warm pipette, quickly apply a drop of 2% liquid agar to the bone marrow cylinder. 
	&#8203;NOTE: The agar solidifies quickly while cooling. To ensure a homogenous covering of the bone marrow, the agar solution has to be kept warm until it is deposited onto the slide.</li>
	<li>Quickly place the slide rapidly on ice until the agar solidifies (1-2 min).</li>
	<li>Under a microscope, use a sharp razor blade to cut and discard the extremities of the bone marrow cylinder because of possible tissue compression in these areas. Transfer the marrow blocks in 1.5 mL microcentrifuge tubes containing 1 mL of cacodylate buffer.</li>
</ol>
3. Embedding bone marrow in resin
CAUTION: Resin components are toxic; some are carcinogenic and must be handled with care under a chemical extraction hood. Use appropriate protective equipment such as gloves and protection glasses. Osmium tetroxide is highly volatile at room temperature and its vapors are very harmful to the eyes, nose, and throat. Before being discarded, 2% osmium tetroxide must be neutralized by adding twice its volume of vegetable oil.
<ol>
	<li>Prepare the epoxy resin as described in the Supplementary File.</li>
	<li>Resin embedding 
	NOTE: Keep the samples in the same microcentrifuge tubes during incubations in successive baths of osmium, uranyl acetate and ethanol. Aspirate the supernatants with a Pasteur pipette. The volume of solution used for each bath must equal at least 10x the volume of the sample.
	<ol>
		<li>Post-fix the blocks with 1% osmium in cacodylate buffer for 1 h at 4 &#176;C, wash once in cacodylate buffer and then once in distilled water.</li>
		<li>Stain with 4% uranyl acetate in distilled water for 1 h, wash twice in distilled water.</li>
		<li>Dehydrate through a graded series of ethanol in distilled water: 4 times in 75% ethanol for 5 min, followed by 3 times in 95% ethanol for 20 min and then 3 times in 100% ethanol for 20 min. At this step, take one syringe of epoxy resin out from the freezer. 
		NOTE: The protocol can be paused in 100% ethanol for 1 h.</li>
		<li>To obtain uniform infiltration and polymerization of epoxy resin inside the marrow, incubate first the blocks in 2 successive baths of propylene oxide for 15 min.</li>
		<li>Add a 1:1 mixture of 100% propylene oxide and epoxy resin and incubate for 1 h. Place the samples on a slow rotary shaker at room temperature.</li>
		<li>Add 100% epoxy resin leave the sample for overnight incubation under agitation.</li>
		<li>Add 100% epoxy resin for 2 h incubation, still under agitation.</li>
		<li>Under a microscope, place the marrow blocks into flat silicone molds. Orientate samples to permit subsequent transversal sectioning of the entire bone marrow. Fill the molds with epoxy resin and place them at 60 &#176;C for 48 h. 
		&#8203;NOTE: All solutions (except ethanol and propylene oxide) are filtered through 0.22 &#181;m filter to avoid samples contamination. To ensure adequate polymerization of the resin, avoid bubbles while filling the molds.</li>
	</ol>
	</li>
</ol>
4. Ultrathin sectioning (Figure 1B)
NOTE: Transmission EM requires thin tissue sections through which electrons can pass generating a projection image of the interior of cells, structure, and organization of inner organelles (granules, endoplasmic reticulum, Golgi) and the arrangement of intracellular cell membranes.
<ol>
	<li>Mount the sample block in an ultra-microtome support. Put it on the sample holder. Trim the samples at 45&#730; in order to remove the excess of resin around the tissue with a rotating diamond or tungsten milling cutter.</li>
	<li>Mount the samples on the ultramicrotome with a diamond knife blade equipped with a water tank. Cut transverse sections of 500 nm and 100 nm thickness for histological and TEM analyses, respectively. Collect floating sections on the water-surface with a loop.</li>
	<li>Deposit the 500 nm thick section on a glass slide and 100 nm thick sections on 200 mesh thin-bar copper grids with a paper filter underneath. Prepare five grids for one condition: stain two grids first and keep the three remaining grids as a backup if necessary.</li>
</ol>
5. Toluidine blue staining for histology
NOTE: Staining sections for histology is important for three reasons: 1) to make sure that the tissue is actually cut and not the resin, 2) to check the quality of the inclusion, and 3) to rapidly evaluate the marrow sample. If this is not correct, cut deeper in the block.
<ol>
	<li>Dry the semi-thin sections slide on a heat plate (60 &#176;C).</li>
	<li>Add filtered 1% toluidine blue/1 % sodium borate in distilled water on the slides and heat on a hot plate (60 &#176;C) for 1-2 min. Wash the slides with distilled water and let it dry on the heat plate.</li>
	<li>Mount sections on coverslips with a drop of Poly(butyl methacrylate-co-methyl methacrylate) mounting medium and examine under a light microscope.</li>
</ol>
6. Heavy metal staining for TEM observation (Figure 1C)
NOTE: For the contrast, the upper side of the grids are inverted on 100 &#181;L drops of each successive bath with a loop. Prior to use, each solution is 0.22 &#181;m filtered. Remove the excess of liquid between each bath by gently contact the grid side on a filter paper.
<ol>
	<li>Stain with 4% uranyl acetate in distilled water for 5 min.</li>
	<li>Wash 3 times in distilled water for 5 min.</li>
	<li>Stain with lead citrate for 3 min.</li>
	<li>Wash 3 times in distilled water for 5 min.</li>
	<li>Deposit the grids by the lower side in contact with the filter paper to let them dry. 
	&#8203;NOTE: Heavy metals react in the presence of carbon dioxide. To minimize the precipitates, avoid air displacement during the contrast, do not speak, keep the environment calm and turn off the air-conditioning.</li>
</ol>
7. TEM (Figure 1E)
NOTE: The sections are introduced in a TEM microscope and examined at 120 kV.
<ol>
	<li>First examine the sections at low magnification (&#60; 500x) to appreciate the general aspect of the preparation (absence of hole in the resin, folds/compression in the sections, precipitates due to staining).</li>
	<li>Then examine the sections at higher magnification (~ 2000x allowing to distinguish the stage of maturation). Count manually the megakaryocytes from each stage of maturation over whole transversal sections (see Representative Results on how do visually identify each stage). 
	NOTE: Each square of the grids is defined as an area for examination (which equals 16000 &#181;m2 for 200 mesh copper grids).</li>
	<li>To assess the number of megakaryocytes, quantify only the squares that are fully covered with a section. To do so, use a model based on the screening of ranges. Observe a first range of squares from an extremity of the section to another, then another range in the same way, etc. Using this procedure, screen fully and systematically the whole marrow transversal section square by square.</li>
	<li>For each square, score the number of Stage I, II or III megakaryocytes. 
	NOTE: Higher magnifications are required to analyze the granules, the DMS organization, the size of cytoplasmic territories and the polylobulated nucleus.</li>
</ol>

<ol>
	<li>Machlus, K. R., Italiano, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+incredible+journey:+From+megakaryocyte+development+to+platelet+formation.">The incredible journey: From megakaryocyte development to platelet formation.</a> The Journal of Cell Biology. 201 (6), 785-796 (2013).</li><li>Zucker-Franklin, D., Termin, C. S., Cooper, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+changes+in+the+megakaryocytes+of+patients+infected+with+the+human+immune+deficiency+virus+(HIV-1).">Structural changes in the megakaryocytes of patients infected with the human immune deficiency virus (HIV-1).</a> American Journal of Pathology. 134 (6), 9 (1989).</li><li>Eckly, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biogenesis+of+the+demarcation+membrane+system+(DMS)+in+megakaryocytes.">Biogenesis of the demarcation membrane system (DMS) in megakaryocytes.</a> Blood. 123 (6), 921-930 (2014).</li><li>Scandola, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+electron+microscopy+to+study+megakaryocytes.">Use of electron microscopy to study megakaryocytes.</a> Platelets. , 1-10 (2020).</li><li>Behnke, O., Forer, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+megakaryocytes+to+platelets:+platelet+morphogenesis+takes+place+in+the+bloodstream.">From megakaryocytes to platelets: platelet morphogenesis takes place in the bloodstream.</a> European Journal of Haematology. 60, 3-23 (2009).</li><li>Eckly, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+megakaryocyte+development+in+the+native+bone+marrow+environment.">Characterization of megakaryocyte development in the native bone marrow environment.</a> Platelets and Megakaryocytes. 788, 175-192 (2012).</li><li>Brown, E., Carlin, L. M., Nerlov, C., Lo Celso, C., Poole, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+membrane+extrusion+sites+drive+megakaryocyte+migration+into+bone+marrow+blood+vessels.">Multiple membrane extrusion sites drive megakaryocyte migration into bone marrow blood vessels.</a> Life Science Alliance. 1 (2), 201800061 (2018).</li><li>Eckly, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Megakaryocytes+use+in+vivo+podosome-like+structures+working+collectively+to+penetrate+the+endothelial+barrier+of+bone+marrow+sinusoids.">Megakaryocytes use in vivo podosome-like structures working collectively to penetrate the endothelial barrier of bone marrow sinusoids.</a> Journal of Thrombosis and Haemostasis. , 15024 (2020).</li><li>Cramer, E. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ultrastructure+of+platelet+formation+by+human+megakaryocytes+cultured+with+the+Mpl+ligand.">Ultrastructure of platelet formation by human megakaryocytes cultured with the Mpl ligand.</a> Blood. 89 (7), 2336-2346 (1997).</li><li>Heijnen, H. F. G., Debili, N., Vainchencker, W., Breton-Gorius, J., Geuze, H. 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The authors have no conflicts of interests to declare.

Direct examination of megakaryocytes in their native environment is essential to understand megakaryopoiesis and platelet formation. In this manuscript, we provide a transmission electron microscopy method combining bone marrow flushing and fixation by immersion, allowing to dissect in situ the morphology characteristics of the entire process of megakaryocyte morphogenesis taking place in the bone marrow.
The flushing of the bone marrow is a critical step of this method, as the succes...

Megakaryocytes are specialized large polyploid cells, localized in the bone marrow, responsible for platelet production1. They originate from hematopoietic stem cells through an intricate maturation process, during which megakaryocyte precursors progressively increase in size, while undergoing extensive concomitant morphologic changes in the cytoplasm and nucleus2. During maturation, megakaryocytes develop a number of distinguishable structural elements including: a polylobulated nucleus, invaginations of the surface membrane that form the demarcation membrane system (DMS), a peripheral zone devoid of organelles surrounded by the actin based cytoskeletal network, and numerous organelles including &#945;-granules, dense granules, lysosomes, and multiple Golgi complexes. At the ultrastructural level, a major modification observed is the cytoplasmic compartmentalization into discrete regions delimited by the DMS3. This extensive supply of membranes will fuel the extension of long cytoplasmic processes in the initial phase of platelet production, which will then remodel into platelets inside the circulation. Any defect during megakaryocyte differentiation and maturation can affect platelet production in term of platelet count and/or platelet function. 
Thin layer transmission electron microscopy (TEM) has been the imaging approach of choice for decades providing high-quality ultrastructure of megakaryocytes that have shaped our understanding of the physiology of thrombopoiesis4,5. This paper focuses on a standardized TEM method allowing to capture the process of platelet biogenesis occurring in situ within the native bone marrow microenvironment, which could also serve as a basis to analyze any bone marrow cell type. We provide ultrastructural examples of the development of megakaryocytes from immature to fully mature, which extend cytoplasmic processes into the microcirculation of sinusoids6. We also describe an easy procedure to quantify the different megakaryocyte maturation stages, instructing on the regeneration and platelet production capacity of the bone marrow.

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			<td>2,4,6-Tri(dimethylaminomethyl)phenol (DMP-30)</td>
			<td>Ladd Research Industries, USA</td>
			<td>21310</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose type LM-3 Low Melting Point Agar</td>
			<td>Electron Microscopy Sciences, USA</td>
			<td>1670-B</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2 Calcium chloride hexahydrate</td>
			<td>Merck, Germany</td>
			<td>2083</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper grids 200 mesh thin-bar</td>
			<td>Oxford Instrument, Agar Scientifics, England</td>
			<td>T200-CU</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethylarsinic acid sodium salt trihydrate</td>
			<td>Merck, Germany</td>
			<td>8.20670.0250</td>
			<td></td>
		</tr>
		<tr>
			<td>Dodecenyl Succinic Anhydride (DDSA)</td>
			<td>Ladd Research Industries, USA</td>
			<td>21340</td>
			<td></td>
		</tr>
		<tr>
			<td>Double Edge Stainless Razor blade</td>
			<td>Electron Microscopy Sciences-EMS, USA</td>
			<td>EM-72000</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol absolut</td>
			<td>VWR International, France</td>
			<td>20821296</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter paper, 90 mm diameter</td>
			<td>Whatman, England</td>
			<td>512-0326</td>
			<td></td>
		</tr>
		<tr>
			<td>Flat embedding silicone mould</td>
			<td>Oxford Instrument, Agar Scientific, England</td>
			<td>G3533</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde 25%</td>
			<td>Electron Microscopy Sciences-EMS, USA</td>
			<td>16210</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat plate Leica EMMP</td>
			<td>Leica Microsystems GmbH, Austria</td>
			<td>705402</td>
			<td></td>
		</tr>
		<tr>
			<td>Histo Diamond Knife 45&#176;</td>
			<td>Diatome, Switzerland</td>
			<td>1044797</td>
			<td></td>
		</tr>
		<tr>
			<td>JEOL 2100 Plus TEM microscope</td>
			<td>JEOL, Japan</td>
			<td>EM-21001BU</td>
			<td></td>
		</tr>
		<tr>
			<td>Lead citrate - Ultrostain 2</td>
			<td>Leica Microsystems GmbH, Austria</td>
			<td>70 55 30 22</td>
			<td></td>
		</tr>
		<tr>
			<td>LX-112 resin</td>
			<td>Ladd Research Industries, USA</td>
			<td>21310</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2 Magnesium chloride hexahydrate</td>
			<td>Sigma, France</td>
			<td>M2393-100g</td>
			<td></td>
		</tr>
		<tr>
			<td>Mounting medium - Poly(butyl methacrylate-co-methyl methacrylate)</td>
			<td>Electron Microscopy Sciences-EMS, USA</td>
			<td>15320</td>
			<td></td>
		</tr>
		<tr>
			<td>Nadic Methyl Anhydride (NMA)</td>
			<td>Ladd Research Industries, USA</td>
			<td>21350</td>
			<td></td>
		</tr>
		<tr>
			<td>Osmium tetroxide 2%</td>
			<td>Merck, Germany</td>
			<td>19172</td>
			<td></td>
		</tr>
		<tr>
			<td>Propylene oxide (1.2-epoxypropane)</td>
			<td>Sigma, France</td>
			<td>82320-250ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Saline injectable solution 0.9% NaCl</td>
			<td>C.D.M Lavoisier, France</td>
			<td>MA 575 420 6</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Surgical steel blade</td>
			<td>Swann-Morton, England</td>
			<td>.0508</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium tetraborate - Borax</td>
			<td>Sigma, France</td>
			<td>B-9876</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Merck, Germany</td>
			<td>84100-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe filter 0.2&#181;m</td>
			<td>Pall Corporation, USA</td>
			<td>514-4126</td>
			<td></td>
		</tr>
		<tr>
			<td>Toluidine blue</td>
			<td>Ladd Research Industries, USA</td>
			<td>N10-70975</td>
			<td></td>
		</tr>
		<tr>
			<td>Trimmer EM TRIM2</td>
			<td>Leica Microsystems GmbH, Austria</td>
			<td>702801</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultramicrotome Ultracut UCT</td>
			<td>Leica Microsystems GmbH, Austria</td>
			<td>656201</td>
			<td></td>
		</tr>
		<tr>
			<td>Uranyl acetate</td>
			<td>Ladd Research Industries, USA</td>
			<td>23620</td>
			<td></td>
		</tr>
	</tbody>
</table>

in situ exploration of murine megakaryopoiesis using transmission electron microscopy

Differentiation and maturation of megakaryocytes occur in close association with the cellular and extracellular components of the bone marrow. These processes are characterized by the gradual appearance of essential structures in the megakaryocyte cytoplasm such as a polyploid and polylobulated nucleus, an internal membrane network called demarcation membrane system (DMS) and the dense and alpha granules that will be found in circulating platelets. In this article, we describe a standardized protocol for the in situ ultrastructural study of murine megakaryocytes using transmission electron microscopy (TEM), allowing for the identification of key characteristics defining their maturation stage and cellular density in the bone marrow. Bone marrows are flushed, fixed, dehydrated in ethanol, embedded in plastic resin, and mounted for generating cross-sections. Semi-thin and thin sections are prepared for histological and TEM observations, respectively. This method can be used for any bone marrow cell, in any EM facility and has the advantage of using small sample sizes allowing for the combination of several imaging approaches on the same mouse.

Bone marrow histology 
Observation of the bone marrow toluidine blue histology under a light microscope is key to rapidly analyze the overall tissue architecture in terms of e.g., tissue compactness, microvessel continuity, and the size and shape of megakaryocytes (Figure 1D). It is performed before ultrathin sections to determine the need of cutting deeper in the bone marrow block. Due to their giant size and nuclear lobulation, the more mature megakar...

Watch this Scientific Journal Video about In Situ Exploration of Murine Megakaryopoiesis using Transmission Electron Microscopy at JoVE.com

In Situ Exploration of Murine Megakaryopoiesis using Transmission Electron Microscopy

Here, we present a protocol to analyze ultrastructure of the megakaryocytes in situ using transmission electron microscopy (TEM). Murine bone marrows are collected, fixed, embedded in epoxy resin and cut in ultrathin sections. After contrast staining, the bone marrow is observed under a TEM microscope at 120 kV.

In Situ Exploration of Murine Megakaryopoiesis using Transmission Electron Microscopy

Differentiation and maturation of megakaryocytes occur in close association with the cellular and extracellular components of the bone marrow. These ...

in-situ-exploration-murine-megakaryopoiesis-using-transmission

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2.5K Views.  Université de Strasbourg, INSERM, EFS Grand Est, BPPS UMR-S 1255, FMTS. Here, we present a protocol to analyze ultrastructure of the megakaryocytes in situ using transmission electron microscopy (TEM). Murine bone marrows are collected, fixed, embedded in epoxy resin and cut in ultrathin sections. After contrast staining, the bone marrow is observed under a TEM microscope at 120 kV. 

Video: In Situ Exploration of Murine Megakaryopoiesis using Transmission Electron Microscopy

In situ Imaging of the Mouse Thymus Using 2-Photon Microscopy

Two-photon Microscopy (TPM) enables us to image deep into the thymus and document the events that are important for thymocyte development. To follow the migration of individuals in a crowd of thymocytes , we generate neonatal chimeras where less than one percent of the thymocytes are derived from a donor that is transgenic for a ubiquitously express fluorescent protein. To generate these partial hematopoetic chimeras, neonatal recipients are injected with bone marrow between 3-7 days of age. After 4-6 weeks, the mouse is sacrificed and the thymus is carefully dissected and bissected preserving the architecture of the tissue that will be imaged. The thymus is glued onto a coverslip in preparation for ex vivo imaging by TPM. During imaging the thymus is kept in DMEM without phenol red that is perfused with 95% oxygen and 5% carbon dioxide and warmed to 37&deg;C. Using this approach, we can study the events required for the generation of a diverse T cell repertoire.

We present step-by-step instructions for the generation of neonatal chimeras as well as the dissection and preparation of the thymus for ex vivo imaging by 2-Photon Microscopy.

Two-photon Microscopy (TPM) enables us to image deep into the thymus and document the events that are important for thymocyte development.  To follow the ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for protein localization, but subcellular context is often absent in fluorescence images. Immuno-electron microscopy, on the other hand, can localize proteins, but the technique is limited by a lack of compatible antibodies, poor preservation of morphology and because most antigens are not exposed to the specimen surface. Correlative approaches can acquire the fluorescence image from a whole cell first, either from immuno-fluorescence or genetically tagged proteins. The sample is then fixed and embedded for electron microscopy, and the images are correlated 1-3. However, the low-resolution fluorescence image and the lack of fiducial markers preclude the precise localization of proteins. 
Alternatively, fluorescence imaging can be done after preserving the specimen in plastic. In this approach, the block is sectioned, and fluorescence images and electron micrographs of the same section are correlated 4-7. However, the diffraction limit of light in the correlated image obscures the locations of individual molecules, and the fluorescence often extends beyond the boundary of the cell. 
Nano-resolution fluorescence electron microscopy (nano-fEM) is designed to localize proteins at nano-scale by imaging the same sections using photo-activated localization microscopy (PALM) and electron microscopy. PALM overcomes the diffraction limit by imaging individual fluorescent proteins and subsequently mapping the centroid of each fluorescent spot 8-10. 
We outline the nano-fEM technique in five steps. First, the sample is fixed and embedded using conditions that preserve the fluorescence of tagged proteins. Second, the resin blocks are sectioned into ultrathin segments (70-80 nm) that are mounted on a cover glass. Third, fluorescence is imaged in these sections using the Zeiss PALM microscope. Fourth, electron dense structures are imaged in these same sections using a scanning electron microscope. Fifth, the fluorescence and electron micrographs are aligned using gold particles as fiducial markers. In summary, the subcellular localization of fluorescently tagged proteins can be determined at nanometer resolution in approximately one week.

We describe a method to localize fluorescently tagged proteins in electron micrographs. Fluorescence is first localized using photo-activated localization microscopy on ultrathin sections. These images are then aligned to electron micrographs of the same section.

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

Plunge Freezing: A Tool for the Ultrastructural and Immunolocalization Studies of Suspension Cells in Transmission Electron Microscopy

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize macromolecular complexes at very high resolution. However, to get as close as possible to the native state, perfect sample preservation is required. Conventional electron microscopy (EM) fixation with aldehydes, for instance, does not provide good ultrastructural preservation. The slow penetration of fixatives induces cell reorganization and loss of various cell components. Therefore, conventional EM fixation does not allow for an instantaneous stabilization and preservation of structures and antigenicity. The best choice for examining intracellular events is to use cryofixation followed by the freeze-substitution fixation method that keeps cells in their native state. High-pressure freezing/freeze-substitution, which preserves the integrity of cellular ultrastructure, is the most commonly used method, but requires expensive equipment. Here, an easy-to-use and low-cost freeze fixation method followed by freeze-substitution for suspension cell cultures is presented.

This manuscript describes an easy-to-use and low-cost cryofixation method for visualizing suspension cells by transmission electron microscopy.

Transmission Electron Microscopy (TEM) is an extraordinary tool for studying cell ultrastructure, in order to localize proteins and visualize ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.