This work was supported by INSERM, Universit&#233; C&#244;te d&#8217;Azur, and by grants from the French National Research Agency (ANR) through the Investments for the Future Labex SIGNALIFE (ANR-11-LABX-0028-01), the program UCA JEDI (ANR-15-IDEX-01) via Academy 2 &#8220;Syst&#232;mes Complexes&#8221; and Academy 4 &#8220;Complexit&#233; et diversit&#233; du vivant&#8221;, Fondation pour la Recherche M&#233;dical (&#201;quipe FRM DEQ20180839587), and the Young Investigator Program to J.G. (ANR18-CE14-0035-01-GILLERON). We also thank the Imaging Core Facility of C3M funded by the Conseil D&#233;partemental des Alpes-Maritimes and the R&#233;gion PACA, and which is also supported by the IBISA Microscopy and Imaging Platform C&#244;te d&#8217;Azur (MICA). We thank Marion Dussot for technical help in tissue preparation. We thank Abby Cuttriss, UCA International Scientific Visibility, for proof reading of the manuscript.

This protocol was tested and is validated for all mouse and human white adipose tissue depots. Human and mouse adipose tissues were collected accordingly to European laws and approved by French and Swedish Ethical committees.
1. Fixation of mouse and human white adipose tissue
<ol>
	<li>Immerse the harvested mouse or human white adipose tissues in at least 10 mL of PBS containing 4% paraformaldehyde (PFA) in a 15 mL plastic tube.</li>
	<li>Shake the plastic tube at room temperature on a rolling plate for 1 h.</li>
	<li>Leave the plastic tube at 4 &#176;C on a rolling plate overnight, to complete the fixation. 
	NOTE: This protocol is adapted for large adipose tissue samples, such as whole epididymal fat pads obtained from mice fed a normal diet (&#8776;250 mg - &#8776;0.6 cm3). For larger samples like epididymal fat pads obtained from mice fed a high fat diet (&#8776;1.5 g - &#8776;4 cm3) or human adipose tissue samples, although the clearing protocol should perfectly work for the whole sample (by scaling-up the PFA and the antibody mixtures), in general, we cut tissue pieces around 1 g (&#8776;2.5 cm3) to reserve the remaining samples either for additional staining or applications. For the PFA (step 1.1.) we recommend using roughly 10 times the volume of the tissue. For the antibody mixtures (steps 3.1. and 3.4.), increase the volume to completely immerse the tissue, and use a 15 mL plastic tube (see Table of Materials) if the tissue is too large for a 1.5 mL plastic microtube (see Table of Materials).</li>
</ol>
2. Permeabilization and saturation of mouse and human white adipose tissue
<ol>
	<li>Rinse the fixed white adipose tissue in 10 mL of PBS for 5 min at room temperature to remove all traces of PFA.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.3% glycine (see Table of Materials) and shake the tube at room temperature in an orbital shaker for 1 h at 100 revolutions per minute (rpm) to quench the remaining free aldehyde groups.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100 (see Table of Materials) and shake the tube at 37 &#176;C in a temperature-controlled orbital shaker for 2 h at 100 rpm.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100 and 20% DMSO (see Table of Materials) and shake the tube at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm overnight.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.1% Tween-20 (see Table of Materials), 0.1% Triton X-100, 0.1% deoxycholate (see Table of Materials) and 20% DMSO and shake the tube at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for at least 24 h.</li>
	<li>Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100 and shake the tube at room temperature in an orbital shaker at 100 rpm for 1 h.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO and 3% BSA (see Table of Materials) and shake the tube at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for 12 h, to saturate any sites that could non-specifically bind antibodies. 
	NOTE: In step 2.7, the BSA can be substituted by blood serum of the species of the secondary antibody used.</li>
</ol>
3. Staining procedure for mouse and human white adipose tissue
<ol>
	<li>Transfer the tissue in a 1.5 mL plastic microtube containing 300 &#181;L of PBS supplemented with 0.2% Triton X-100, 10% DMSO, 3% BSA and the primary antibodies (10x more concentrated than for cryosection staining but optimal antibody concentration should be evaluated for each antibody). Protect the tube from light by covering with aluminium foil and shake the tube at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for at least two&#160;days (see the note in step 1.1 and step 3.7.1).</li>
	<li>Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO and 3% BSA and shake the tube protected from light at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for 5 h. Perform this step twice.</li>
	<li>Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO and 3% BSA and shake the tube, protected from light, at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for one night to two days.</li>
	<li>Transfer the tissue to a 1.5 mL plastic microtube containing 300 &#181;L of PBS supplemented with 0.2% Triton X-100, 10% DMSO, 3% BSA and the secondary antibodies (10x more concentrated than for cryosection staining). Protect the tube from light by covering with aluminium foil and shake the tube at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for at least two&#160;days (see the note in step 1.1 and step 3.7.1).</li>
	<li>Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO, and 3% BSA and shake the tube, protected from light, at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for 5 h. Perform this step twice.</li>
	<li>Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS supplemented with 0.2% Triton X-100, 10% DMSO, and 3% BSA and shake the tube, protected from light, at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for one night to two days.</li>
	<li>Rinse the tissue in a 15 mL plastic tube containing 10 mL of PBS and shake the tube, protected from light, at 37 &#176;C in a temperature-controlled orbital shaker at 100 rpm for 2 h. 
	NOTE: For the labelling of specific structures such as nuclei or actin, add 4&#8242;,6-diamidino-2-phenylindole (DAPI or equivalent) and/or the fluorescently labelled-phalloidin have to be added at step 3.1. when only fluorescently labelled primary antibodies are used, or at step 3.4. when secondary antibodies are used. 
	NOTE: For the staining procedure, primary antibodies already conjugated with fluorochromes (like those used for flow cytometry) can be used and we recommend it for the gain of time and specificity. Indeed, even if mouse primary antibodies can be used followed by secondary anti-mouse antibodies, as we demonstrated it here, we have often observed non-specific staining of blood vessels due to circulating immunoglobulin. Therefore, to avoid this issue, primary antibodies that are already labelled are highly recommended and here we provide evidence that the antibodies used in flow cytometry are compatible with our procedure. However, in the specific case of an antigen that is expressed at low levels, a signal amplification step via secondary antibodies is mandatory; when the only available primary antibody is made in mouse, a cardiac-perfusion of the mice with PBS for at least 5 min at the sacrifice can remove a large proportion of circulating immunoglobulin and thus the non-specific staining.</li>
</ol>
4. Clearing procedure for mouse and human white adipose tissue
<ol>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of 50% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of 70% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of 95% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of 100% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm for 2 h.</li>
	<li>Immerse the tissue in a 15 mL plastic tube containing 10 mL of 100% ethanol and shake the tube, protected from light, at room temperature in an orbital shaker at 100 rpm overnight.</li>
	<li>Immerse the tissue in a 20 mL glass bottle with a plastic cap (see table of materials) containing 5 mL of methyl salicylate (see Table of Materials) under a chemical hood and shake the glass container, protected from light, at room temperature in an orbital shaker at 100 rpm for at least 2 h. 
	NOTE: The clearing procedure could be significantly accelerated (if necessary) by avoiding steps 4.3. and 4.5., although the final clearing quality would be slightly reduced.</li>
</ol>
5. 3D-confocal imaging of cleared white adipose tissue
<ol>
	<li>Transfer the tissue to a metallic imaging chamber equipped with a glass bottom (see Table of Materials) under a chemical hood and fill the chamber with fresh methyl salicylate. 
	NOTE:&#160;To secure the tissue in place, and thus to prevent it from floating or moving sideways in the chamber, apply multiple 18 mm round glass coverslips (see Table of Materials) on top of the tissue when mounting it into the chamber.</li>
	<li>Place the imaging chamber on an inverted confocal microscope.</li>
	<li>Image the tissue using a low magnification objective (e.g., 4x objective) to generate a few cm3 3D maps of the whole adipose tissue or of the human tissue sample.</li>
	<li>Select several areas for the tissue sampling at higher magnification. Typically, use a 20x long distance air objective that provides a good ratio between resolution and depth. We acquire large mosaic images with z-stacks between 600 and 2000 &#181;m depth. 
	NOTE:&#160;Use a long-distance objective to image deeper into the tissue.</li>
</ol>
6. Extraction of quantitative results from the 3D adipose tissue images
NOTE: The segmentation of the different structures and the subsequent extraction of the quantitative information from the 3D-image stack generated in point 5 can be performed using any of the many existing image analysis software options, either commercial or freeware. In the following points, we describe a strategy that is routinely used in our laboratory to extract quantitative information from 3D adipose tissue images using commercial software (see Table of Materials).
<ol>
	<li>Convert the 3D stacks onto the software format to free-up memory space.</li>
	<li>Segment the cells.
	<ol>
		<li>Open the Cell module of the software.</li>
		<li>Change the Cell Detection setting to plasma membrane staining.</li>
		<li>Choose the fluorescent channel of the marker used to delineate the cell periphery (phalloidin which labels cortical actin or plasma membrane markers such as F4/80 for macrophage, TCR-&#946; for T cells, or cluster of differentiation CD proteins specific for immune cell subtypes).</li>
		<li>Set up the thresholds and provide a range of expected cell size (i.e., 1 to 200 &#181;m for cell detection).</li>
		<li>Run the segmentation. A volume corresponding to the z-stack will be generated with the segmented cells color-coded with a different color for each of the neighbouring cells.</li>
		<li>Apply statistical filters (size, roundness, circularity, etc.) in the statistic tab to exclude segmentation artefacts and/or to refine the segmented cells to the cell of interest based on their size (i.e., 20-200 &#181;m for adipocytes; 5-25 &#181;m for macrophages; 1-3 &#181;m for lymphocytes).</li>
		<li>Extract measurements and quantitative data (volume, number, size, diameter, etc.) from the statistics tab.</li>
	</ol>
	</li>
	<li>Segment cellular components (nuclei, vesicles, etc.) or vessels as a structure. 
	NOTE: Although the filament segmentation is very useful to study the connectivity of the vessels, we use the surface module segmentation to extract data on the size, volume and diameters of the vessels. Indeed, the quantification of the sizes of the vessels is lost when using the filaments module.
	<ol>
		<li>Open the Surface module of the software.</li>
		<li>Select the fluorescent channel of the marker used to specifically label the subcellular component to reconstruct it in 3D.</li>
		<li>Set up the thresholds and provide a range of the expected diameter size of the structure (i.e., 0.5-5 &#181;m for nuclei; 0-100 &#181;m for vessels; etc.).</li>
		<li>Run the segmentation. A volume with the segmented cellular structure will be generated. Measurements and quantitative data (volume, number, size, diameter, etc.) can be extracted from the statistics tab.</li>
	</ol>
	</li>
	<li>Segment the vessels as a tubular continuum. 
	NOTE: Although the filament segmentation is very useful to study the connectivity of the vessels, we use the surface module segmentation to extract data on the size, volume and diameters of the vessels. Indeed, the quantification of the sizes of the vessels is lost when using the filaments module.
	<ol>
		<li>Open the Filaments module of the software.</li>
		<li>Select the fluorescent channel corresponding to the vessel staining.</li>
		<li>Set up the thresholds for the fluorescence intensity and select the appropriate number of expected connecting nodes.</li>
		<li>Run the segmentation. Filaments representing vessel network will be generated in 3D. Measurements can be extracted from the statistics tab, allowing to obtain quantitative data including vessel length, number of vessel branching, etc.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Pellegrinelli, V., Carobbio, S., Vidal-Puig, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose+tissue+plasticity:+how+fat+depots+respond+differently+to+pathophysiological+cues.">Adipose tissue plasticity: how fat depots respond differently to pathophysiological cues.</a> Diabetologia. 59 (6), 1075-1088 (2016).</li><li>Stern, J. H., Rutkowski, J. M., Scherer, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adiponectin,+Leptin,+and+Fatty+Acids+in+the+Maintenance+of+Metabolic+Homeostasis+through+Adipose+Tissue+Crosstalk.">Adiponectin, Leptin, and Fatty Acids in the Maintenance of Metabolic Homeostasis through Adipose Tissue Crosstalk.</a> Cell Metabolism. 23 (5), 770-784 (2016).</li><li>Mittendorfer, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Origins+of+metabolic+complications+in+obesity:+adipose+tissue+and+free+fatty+acid+trafficking.">Origins of metabolic complications in obesity: adipose tissue and free fatty acid trafficking.</a> Current Opinion in Clinical Nutrition &amp; Metabolic Care. 14 (6), 535-541 (2011).</li><li>Hammarstedt, A., Gogg, S., Hedjazifar, S., Nerstedt, A., Smith, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impaired+Adipogenesis+and+Dysfunctional+Adipose+Tissue+in+Human+Hypertrophic+Obesity.">Impaired Adipogenesis and Dysfunctional Adipose Tissue in Human Hypertrophic Obesity.</a> Physiological Reviews. 98 (4), 1911-1941 (2018).</li><li>Moreno-Indias, I., Tinahones, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impaired+adipose+tissue+expandability+and+lipogenic+capacities+as+ones+of+the+main+causes+of+metabolic+disorders.">Impaired adipose tissue expandability and lipogenic capacities as ones of the main causes of metabolic disorders.</a> Journal of Diabetes Research. 2015, 970375 (2015).</li><li>Vergoni, B., 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=DNA+Damage+and+the+Activation+of+the+p53+Pathway+Mediate+Alterations+in+Metabolic+and+Secretory+Functions+of+Adipocytes.">DNA Damage and the Activation of the p53 Pathway Mediate Alterations in Metabolic and Secretory Functions of Adipocytes.</a> Diabetes. 65 (10), 3062-3074 (2016).</li><li>Xue, Y., Xu, X., Zhang, X. Q., Farokhzad, O. C., Langer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preventing+diet-induced+obesity+in+mice+by+adipose+tissue+transformation+and+angiogenesis+using+targeted+nanoparticles.">Preventing diet-induced obesity in mice by adipose tissue transformation and angiogenesis using targeted nanoparticles.</a> Proceedings of the National Academy of Sciences U S A. 113 (20), 5552-5557 (2016).</li><li>Zwick, R. K., 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=Adipocyte+hypertrophy+and+lipid+dynamics+underlie+mammary+gland+remodeling+after+lactation.">Adipocyte hypertrophy and lipid dynamics underlie mammary gland remodeling after lactation.</a> Nature Communication. 9 (1), 3592 (2018).</li><li>Zhang, L., 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=The+inflammatory+changes+of+adipose+tissue+in+late+pregnant+mice.">The inflammatory changes of adipose tissue in late pregnant mice.</a> Journal of Molecular Endocrinology. 47 (2), 157-165 (2011).</li><li>Yang, H., 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=Obesity+increases+the+production+of+proinflammatory+mediators+from+adipose+tissue+T+cells+and+compromises+TCR+repertoire+diversity:+implications+for+systemic+inflammation+and+insulin+resistance.">Obesity increases the production of proinflammatory mediators from adipose tissue T cells and compromises TCR repertoire diversity: implications for systemic inflammation and insulin resistance.</a> Journal of Immunology. 185 (3), 1836-1845 (2010).</li><li>Laforest, S., 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=Comparative+analysis+of+three+human+adipocyte+size+measurement+methods+and+their+relevance+for+cardiometabolic+risk.">Comparative analysis of three human adipocyte size measurement methods and their relevance for cardiometabolic risk.</a> Obesity. 25 (1), 122-131 (2017).</li><li>Azaripour, 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=A+survey+of+clearing+techniques+for+3D+imaging+of+tissues+with+special+reference+to+connective+tissue.">A survey of clearing techniques for 3D imaging of tissues with special reference to connective tissue.</a> Progress in Histochemistry and Cytochemistry. 51 (2), 9-23 (2016).</li><li>Richardson, D. S., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clarifying+Tissue+Clearing.">Clarifying Tissue Clearing.</a> Cell. 162 (2), 246-257 (2015).</li><li>Chi, J., Crane, A., Wu, Z., Cohen, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipo-Clear:+A+Tissue+Clearing+Method+for+Three-Dimensional+Imaging+of+Adipose+Tissue.">Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue.</a> Journal of Visualized Experiments. (137), e58271 (2018).</li><li>Roberts, D. G., Johnsonbaugh, H. B., Spence, R. D., MacKenzie-Graham, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+Clearing+of+the+Mouse+Central+Nervous+System+Using+Passive+CLARITY.">Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY.</a> Journal of Visualized Experiments. (112), e540225 (2016).</li><li>Woo, J., Lee, M., Seo, J. M., Park, H. S., Cho, Y. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+the+optical+transparency+of+rodent+tissues+by+modified+PACT-based+passive+clearing.">Optimization of the optical transparency of rodent tissues by modified PACT-based passive clearing.</a> Experimental &amp; Molecular Medicine. 48 (12), 274 (2016).</li><li>Zhang, Y., 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=3D+imaging+of+optically+cleared+tissue+using+a+simplified+CLARITY+method+and+on-chip+microscopy.">3D imaging of optically cleared tissue using a simplified CLARITY method and on-chip microscopy.</a> Science Advances. 3 (8), 1700553 (2017).</li><li>Ke, M. T., Imai, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+clearing+of+fixed+brain+samples+using+SeeDB.">Optical clearing of fixed brain samples using SeeDB.</a> Current Protocols in Neuroscience. 66, 22 (2014).</li><li>Hahn, 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=High-resolution+imaging+of+fluorescent+whole+mouse+brains+using+stabilised+organic+media+(sDISCO).">High-resolution imaging of fluorescent whole mouse brains using stabilised organic media (sDISCO).</a> Journal of Biophotonics. 12 (8), 201800368 (2019).</li><li>Spalteholz, W., Hierzel, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> &#220;ber das Durchsichtigmachen von Menchlichen und Tierichen Pr&#228;paraten und Seine Theoretischen. , (1911).</li><li>Shields, K. J., Wu, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+Adipose+Tissue+Proteomics.">Differential Adipose Tissue Proteomics.</a> Methods in Molecular Biology. 1788, 243-250 (2018).</li><li>Bourlier, V., 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=Remodeling+phenotype+of+human+subcutaneous+adipose+tissue+macrophages.">Remodeling phenotype of human subcutaneous adipose tissue macrophages.</a> Circulation. 117 (6), 806-815 (2008).</li><li>Hagberg, C. E., 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=Flow+Cytometry+of+Mouse+and+Human+Adipocytes+for+the+Analysis+of+Browning+and+Cellular+Heterogeneity.">Flow Cytometry of Mouse and Human Adipocytes for the Analysis of Browning and Cellular Heterogeneity.</a> Cell Reports. 24 (10), 2746-2756 (2018).</li><li>Hill, D. 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=Distinct+macrophage+populations+direct+inflammatory+versus+physiological+changes+in+adipose+tissue.">Distinct macrophage populations direct inflammatory versus physiological changes in adipose tissue.</a> Proceedings of the National Academy of Sciences U S A. 115 (22), 5096-5105 (2018).</li><li>Acosta, J. R., 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=Single+cell+transcriptomics+suggest+that+human+adipocyte+progenitor+cells+constitute+a+homogeneous+cell+population.">Single cell transcriptomics suggest that human adipocyte progenitor cells constitute a homogeneous cell population.</a> Stem Cell Research &amp; Therapy. 8 (1), 250 (2017).</li><li>Coppack, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose+tissue+changes+in+obesity.">Adipose tissue changes in obesity.</a> Biochemical Society Transactions. 33, 1049-1052 (2005).</li><li>Cancello, R., 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=Reduction+of+macrophage+infiltration+and+chemoattractant+gene+expression+changes+in+white+adipose+tissue+of+morbidly+obese+subjects+after+surgery-induced+weight+loss.">Reduction of macrophage infiltration and chemoattractant gene expression changes in white adipose tissue of morbidly obese subjects after surgery-induced weight loss.</a> Diabetes. 54 (8), 2277-2286 (2005).</li><li>Cinti, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipocyte+differentiation+and+transdifferentiation:+plasticity+of+the+adipose+organ.">Adipocyte differentiation and transdifferentiation: plasticity of the adipose organ.</a> Journal of Endocrinology Investigation. 25 (10), 823-835 (2002).</li><li>Wellen, K. E., Hotamisligil, G. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Obesity-induced+inflammatory+changes+in+adipose+tissue.">Obesity-induced inflammatory changes in adipose tissue.</a> Journal of Clinical Investigation. 112 (12), 1785-1788 (2003).</li><li>Li, X., Mao, Z., Yang, L., Sun, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Co-staining+Blood+Vessels+and+Nerve+Fibers+in+Adipose+Tissue.">Co-staining Blood Vessels and Nerve Fibers in Adipose Tissue.</a> Journal of Visualized Experiments. (144), e59266 (2019).</li><li>Gustafsson, N., 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=Fast+live-cell+conventional+fluorophore+nanoscopy+with+ImageJ+through+super-resolution+radial+fluctuations.">Fast live-cell conventional fluorophore nanoscopy with ImageJ through super-resolution radial fluctuations.</a> Nature Communication. 7, 12471 (2016).</li></ol>

The authors have no conflicts to disclose.

The modifications that occur within the adipose tissue over the course of pathological progression, such as that of obesity, is fundamental to the understanding of the mechanisms behind the pathology. Pioneering studies that revealed such mechanisms in adipose tissue have been based on global approaches such as whole adipose tissue proteomics21, flow cytometry22,23, and transcriptomics24,<sup class="xref"...

Obesity is characterized by an increase in adipose tissue mass and has become a major worldwide public health issue, given that people with obesity have increased risk of developing cardiovascular disease, type-2 diabetes, liver diseases and some cancers.
A fundamental physiological function of adipose tissue is to modulate whole-body glucose and lipid homeostasis1,2. During the feeding period, the adipocytes (i.e., the main cells of the adipose tissue) store&#160;the excess of glucose and lipids provided by a meal into triglycerides. During fasting, the adipocytes break down the triglycerides into non-esterified fatty acids and glycerol to sustain the energy demand of the body. During the development of obesity, adipose tissue expand by increasing the size (hypertrophia) and/or the number (hyperplasia) of adipocytes1, to increase their storage capacity. When the expansion of adipose tissue reaches its limit, a constant highly variable among patients, the remaining lipids accumulates into other metabolic organs including muscles and liver3,4, leading to their functional failure and initiating obesity-related cardio-metabolic complications1,5. Therefore, identifying the mechanisms that govern adipose tissue expansion is a key clinical challenge.
The morphological modifications documented within adipose tissues during obesity are linked to its pathological dysfunction. Several staining procedures have been used to describe the tissue organization of the adipose tissue, including actin6, vascular markers7, lipid-droplet markers8, and specific immune cell markers9,10. However, because of the huge diameter of adipocytes (50 to 200 &#181;m)11, it is essential to analyze a large portion of the whole tissue in three dimensions in order to accurately analyze the dramatic structural AT changes observed during obesity. However, because the light does not penetrate an opaque tissue, imaging in 3D within a large tissue samples using fluorescence microscopy is not possible. Methods of tissue clearing to make them transparent have been reported in the literature (for a review, see12) allowing one to clear tissues and to perform in-depth, whole tissue fluorescence microscopy. These methods offer unprecedented opportunities to assess the 3D cellular organization in healthy and diseased tissue. Each of the described methods have advantages and drawbacks, and therefore need to be carefully selected depending on the studied tissue (for a review, see13). Indeed, some approaches require a long incubation period and/or the use of materials or compounds that are either expensive, toxic or difficult to obtain14,15,16,17,18,19. Taking advantage of one of the first compounds used a century ago by Werner Spalteholz to clear tissues20, we set up a user-friendly and inexpensive protocol that is very well adapted for the clearing of all mouse and human adipose tissue depots in any laboratory with typical equipment including a chemical hood, a temperature-controlled orbital shaker and a confocal microscope.

<table>
	<tbody>
		<tr>
			<td>1.5 mL microtubes</td>
			<td>Eppendorff tubes - Dutscher</td>
			<td>33528</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL plastic tubes</td>
			<td>Falcon tubes - Dutscher</td>
			<td>352096</td>
			<td></td>
		</tr>
		<tr>
			<td>18 mm round glass coverslip</td>
			<td>Mariendfeld</td>
			<td>0117580</td>
			<td></td>
		</tr>
		<tr>
			<td>20 mL glass bottle</td>
			<td>Wheaton</td>
			<td>986546</td>
			<td></td>
		</tr>
		<tr>
			<td>anti-mouse-alexa647-conjugated antibody</td>
			<td>Jackson ImmunoResearch</td>
			<td>715-605-150</td>
			<td>Dilution: 1/100</td>
		</tr>
		<tr>
			<td>anti-rabbit-alexa647-conjugated antibody</td>
			<td>Jackson ImmunoResearch</td>
			<td>711-605-152</td>
			<td>Dilution: 1/100</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma-aldrich</td>
			<td>A6003</td>
			<td></td>
		</tr>
		<tr>
			<td>CD301-PE antibody</td>
			<td>Biolegend</td>
			<td>BLE145703</td>
			<td>Dilution: 1/100</td>
		</tr>
		<tr>
			<td>CD31 antibody</td>
			<td>AbCam</td>
			<td>ab215912</td>
			<td>Dilution: 1/50</td>
		</tr>
		<tr>
			<td>Commercial 3D analysis software - IMARIS</td>
			<td>Oxford instrument</td>
			<td></td>
			<td>with Cell module</td>
		</tr>
		<tr>
			<td>Confocal microscope - Nikon A1R</td>
			<td>Nikon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dapi</td>
			<td>ThermoFisher</td>
			<td>D1306</td>
			<td>Stock Concentration: 5 mg/mL; dilution 1/1000</td>
		</tr>
		<tr>
			<td>Deoxycholate</td>
			<td>Sigma-aldrich</td>
			<td>D6750</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma-aldrich</td>
			<td>D8418</td>
			<td></td>
		</tr>
		<tr>
			<td>Glut4 antibody</td>
			<td>Santa Cruz</td>
			<td>sc-53566</td>
			<td>Dilution: 1/50</td>
		</tr>
		<tr>
			<td>Glycine</td>
			<td>Sigma-aldrich</td>
			<td>G7126</td>
			<td></td>
		</tr>
		<tr>
			<td>Lectin-DyLight649</td>
			<td>Vector Lab</td>
			<td>DL-1178-1</td>
			<td>Stock Concentration : 2 &#181;g/&#181;L; IV Injection: 50 &#181;L/mice</td>
		</tr>
		<tr>
			<td>Metallic imaging chamber equipped with glass bottom - AttoFluor Chamber</td>
			<td>Thermofisher</td>
			<td>A7816</td>
			<td></td>
		</tr>
		<tr>
			<td>Methyl salicylate</td>
			<td>Sigma-aldrich</td>
			<td>M6752</td>
			<td></td>
		</tr>
		<tr>
			<td>Perilipin antibody</td>
			<td>Progen</td>
			<td>651156</td>
			<td>Dilution: 1/50</td>
		</tr>
		<tr>
			<td>Phalloidin-alexa488</td>
			<td>ThermoFisher</td>
			<td>A12379</td>
			<td>Dilution: 1/100</td>
		</tr>
		<tr>
			<td>TCR-&#946;-PB antibody</td>
			<td>Biolegend</td>
			<td>BLE109225</td>
			<td>Dilution: 1/100</td>
		</tr>
		<tr>
			<td>TH antibody</td>
			<td>AbCam</td>
			<td>ab112</td>
			<td>Dilution: 1/50</td>
		</tr>
		<tr>
			<td>Triton X100</td>
			<td>Sigma-aldrich</td>
			<td>X100</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Sigma-aldrich</td>
			<td>P416</td>
			<td></td>
		</tr>
	</tbody>
</table>

exploring adipose tissue structure by methylsalicylate clearing and 3d imaging

Obesity is a major worldwide public health issue that increases the risk to develop cardiovascular diseases, type-2 diabetes, and liver diseases. Obesity is characterized by an increase in adipose tissue (AT) mass due to adipocyte hyperplasia and/or hypertrophia, leading to profound remodeling of its three-dimensional structure. Indeed, the maximal capacity of AT to expand during obesity is pivotal to the development of obesity-associated pathologies. This AT expansion is an important homeostatic mechanism to enable adaptation to an excess of energy intake and to avoid deleterious lipid spillover to other metabolic organs, such as muscle and liver. Therefore, understanding the structural remodeling that leads to the failure of AT expansion is a fundamental question with high clinical applicability. In this article, we describe a simple and fast clearing method that is routinely used in our laboratory to explore the morphology of mouse and human white adipose tissue by fluorescent imaging. This optimized AT clearing method is easily performed in any standard laboratory equipped with a chemical hood, a temperature-controlled orbital shaker and a fluorescent microscope. Moreover, the chemical compounds used are readily available. Importantly, this method allows one to resolve the 3D AT structure by staining various markers to specifically visualize the adipocytes, the neuronal and vascular networks, and the innate and adaptive immune cells distribution.

Using the procedure described here and summarized in Figure 1, we were able to stain and optically clear human and mouse white adipose tissue as presented in Figure 2A and Figure 2B, respectively. The cleared tissue was transferred to the metallic imaging chamber to perform confocal imaging (Figure 3A). The clearing drastically improved the depth of the tissue images that we were able to acquire (<stron...

Watch this Scientific Journal Video about Exploring Adipose Tissue Structure by Methylsalicylate Clearing and 3D Imaging at JoVE.com

Exploring Adipose Tissue Structure by Methylsalicylate Clearing and 3D Imaging

Here, we describe a simple, inexpensive and fast clearing method to resolve the 3D structure of both mouse and human white adipose tissue using a combination of markers to visualize vasculature, nuclei, immune cells, neurons, and lipid-droplet coat proteins by fluorescent imaging.

Obesity is a major worldwide public health issue that increases the risk to develop cardiovascular diseases, type-2 diabetes, and liver diseases. Obesity ...

exploring-adipose-tissue-structure-methylsalicylate-clearing-3d

Research

JoVE Journal

Biology

7.3K Views.  Inserm UMR1065, C3M, Team "Cellular and Molecular Pathophysiology of Obesity", Nice, France. Here, we describe a simple, inexpensive and fast clearing method to resolve the 3D structure of both mouse and human white adipose tissue using a combination of markers to visualize vasculature, nuclei, immune cells, neurons, and lipid-droplet coat proteins by fluorescent imaging.

Video: Exploring Adipose Tissue Structure by Methylsalicylate Clearing and 3D Imaging

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

ACT-PRESTO: Biological Tissue Clearing and Immunolabeling Methods for Volume Imaging

The identification and exploration of the detailed organization of organs or of the whole body at the cellular level are fundamental challenges in biology. Transitional methods require a substantial amount of time and effort to obtain a 3D image and including sectioning the intact tissue, immunolabeling, and imaging serially-sectioned tissue, which produces a loss of information at each step of the process. In recently developed approaches for high-resolution imaging within intact tissue, molecular characterization has been restricted to the labeling of proteins. However, currently available protocols for organ clearing require a considerably long process time, making it difficult to implement tissue clearing techniques in the lab. We recently established a rapid and highly-reproducible protocol termed ACT-PRESTO (active clarity technique&#8211;pressure related efficient and stable transfer of macromolecules into organs), which allows for tissue clearance within several hours. Moreover, ACT-PRESTO enables rapid immunolabeling with conventional methods and accelerates antibody penetration into the deep layer of densely-formed, thick specimens by applying pressure or convection flow. We describe how to prepare tissues, how to clear by lipid removal using electrophoresis, and how to immuno-stain by a pressure-assisted delivery. The rapidity and consistency of the protocol will expedite the performance of 3D histological research and volume-based diagnoses.

ACT-PRESTO (active clarity technique&#8211;pressure related efficient and stable transfer of macromolecules into organs) enables rapid, efficient, but inexpensive tissue clearing and rapid antibody penetration into cleared tissues by passive diffusion or pressure-assisted delivery. Using this method, a wide range of tissues can be cleared, immunolabeled by multiple antibodies, and volume-imaged.

The identification and exploration of the detailed organization of organs or of the whole body at the cellular level are fundamental challenges in ...

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 ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...

Visualization of 3D White Adipose Tissue Structure Using Whole-mount Staining

Adipose tissue is an important metabolic organ with high plasticity and is responsive to environmental stimuli and nutrient status. As such, various techniques have been developed to study the morphology and biology of adipose tissue. However, conventional visualization methods are limited to studying the tissue in 2D sections, failing to capture the 3D architecture of the whole organ. Here we present whole-mount staining, an immunohistochemistry method that preserves intact adipose tissue morphology with minimal processing steps. Hence, the structures of adipocytes and other cellular components are maintained without distortion, achieving the most representative 3D visualization of the tissue. In addition, whole-mount staining can be combined with lineage tracing methods to determine cell fate decisions. However, this technique has some limitations to providing accurate information regarding deeper parts of adipose tissue. To overcome this limitation, whole-mount staining can be further combined with tissue clearing techniques to remove the opaqueness of tissue and allow for complete visualization of entire adipose tissue anatomy using light-sheet fluorescent microscopy. Therefore, a higher resolution and more accurate representation of adipose tissue structures can be captured with the combination of these techniques.

The focus of the present study is to demonstrate the whole-mount immunostaining and visualization technique as an ideal method for 3D imaging of adipose tissue architecture and cellular component.

Adipose tissue is an important metabolic organ with high plasticity and is responsive to environmental stimuli and nutrient status. As such, various ...

Human Adipose Tissue Micro-fragmentation for Cell Phenotyping and Secretome Characterization

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. Accordingly, techniques for harvesting and processing human adipose tissue have evolved in order to quickly and efficiently obtain large amounts of tissue. Among these, the closed system technology represents an innovative and easy-to-use system to harvest, process, and re-inject refined fat tissue in a short time and in the same intervention (intra-operatively). Adipose tissue is collected by liposuction, washed, emulsified, rinsed and minced mechanically into cell clusters of 0.3 to 0.8 mm. Autologous transplantation of mechanically fragmented adipose tissue has shown remarkable efficacy in different therapeutic indications such as aesthetic medicine and surgery, orthopedic and general surgery. Characterization of micro-fragmented adipose tissue revealed the presence of intact small vessels within the adipocyte clusters; hence, the perivascular niche is left unperturbed. These clusters are enriched in perivascular cells (i.e., mesenchymal stem cell (MSC) ancestors) and in vitro analysis showed an increased release of growth factors and cytokines involved in tissue repair and regeneration, compared to enzymatically derived MSCs. This suggests that the superior therapeutic potential of microfragmented adipose tissue is explained by a higher frequency of presumptive MSCs and enhanced secretory activity. Whether these added pericytes directly contribute to higher growth factor and cytokine production is not known. This clinically approved procedure allows the transplantation of presumptive MSCs without the need for expansion and/or enzymatic treatment, thus bypassing the requirements of GMP guidelines, and reducing the costs for cell-based therapies.

Here, we present human adipose tissue enzyme-free micro-fragmentation using a closed system device. This new method allows the obtainment of sub-millimeter clusters of adipose tissue suitable for in vivo transplantation, in vitro culture, and further cell isolation and characterization.

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. ...

Optical Clearing of Plant Tissues for Fluorescence Imaging

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In addition, autofluorescence attributed to chlorophyll hampers the observation of fluorescent proteins in plants. For a long time, various clearing reagents have been used to make plants transparent. However, conventional clearing reagents diminish fluorescent signals; therefore, it has not been possible to observe the cellular and intracellular structures with fluorescent proteins. Reagents were developed that can clear plant tissues by removing chlorophyll while maintaining fluorescent protein stability. A detailed protocol is provided here for the optical clearing of plant tissues using clearing reagents, ClearSee (CS) or ClearSeeAlpha (CSA). The preparation of cleared plant tissues involves three steps: fixation, washing, and clearing. Fixation is a crucial step in maintaining the cellular structures and intracellular stability of fluorescent proteins. The incubation time for clearing depends on the tissue type and species. In Arabidopsis thaliana, the time required for clearing with CS was 4 days for leaves and roots, 7 days for seedlings, and 1 month for pistils. CS also required a relatively short time of 4 days to make the gametophytic leaves of Physcomitrium patens transparent. In contrast, pistils in tobacco and torenia produced brown pigment due to oxidation during CS treatment. CSA reduced the brown pigment by preventing oxidation and could make tobacco and torenia pistils transparent, although it took a relatively long time (1 or 2 months). CS and CSA were also compatible with staining using chemical dyes, such as DAPI (4',6-diamidino-2-phenylindole) and Hoechst 33342 for DNA and Calcofluor White, SR2200, and Direct Red 23 for the cell wall. This method can be useful for whole-plant imaging to reveal intact morphology, developmental processes, plant-microbe interactions, and nematode infections.

Here, a method is described for making plant tissues transparent while maintaining the stability of fluorescent proteins. This technique facilitates deep imaging of cleared plant tissues without physical sectioning.

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In ...

Differentiation and Imaging of Brown Adipocytes from the Stromal Vascular Fraction of Interscapular Adipose Tissue from Newborn Mice

Brown adipose tissue (BAT) is only present in mammals and has a thermogenic function. Brown adipocytes are characterized by a multilocular cytoplasm with multiple lipid droplets, a central nucleus, a high mitochondrial content, and the expression of uncoupling protein 1 (UCP1). BAT has been proposed as a potential therapeutic target for obesity and its associated metabolic disorders due to its ability to dissipate metabolic energy as heat. To investigate BAT function and regulation, brown adipocyte culturing is indispensable. The present protocol optimizes tissue processing and cell differentiation for culturing brown adipocytes from newborn mice. Additionally, procedures for the imaging of differentiated adipocytes with both confocal immunofluorescence and transmission electron microscopy are shown. In the brown adipocytes differentiated with the techniques described herein, the major defining features of classical BAT are preserved, including high UCP1 levels, increased mitochondrial mass, and very close physical contact between the lipid droplets and mitochondria, making this method a valuable tool for BAT studies.

Preadipocytes are isolated from the stromal vascular fraction of interscapular brown adipose tissue from newborn mice and differentiated into cells that accumulate lipid droplets, express molecular markers, and show the mitochondrial morphology of mature brown adipocytes. These cells are further analyzed by immunofluorescence and transmission electron microscopy.

Brown adipose tissue (BAT) is only present in mammals and has a thermogenic function. Brown adipocytes are characterized by a multilocular cytoplasm with ...