This study was supported by the National Institute of Health (NIH) grant R01DK109001 (to K.S.).

All procedures containing animal subjects have been approved by the Animal Welfare Committee of University of Texas Health Science Center at Houston (animal protocol number: AWC-18-0057).
1. Reagent Preparation
<ol>
	<li>1x phosphate-buffered saline (PBS, pH 7.4): to make 1 L of 1x PBS, dissolve 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, and 0.24 g of KH2PO4 in 800 mL of distilled water. Adjust the pH to 7.4 and fill with distilled water to achieve a final volume of 1 L.</li>
	<li>1% paraformaldehyde in 1x PBS (1% PFA, wt/vol): to achieve a final volume of 50 mL, add 3.125 mL of 16% PFA (see Table of Materials) to 46.875 mL of 1x PBS. Mix well and store at 4 &#176;C for later use. 
	CAUTION: Paraformaldehyde is toxic. All procedures should be carried out under a fume hood to avoid inhalation and skin contact.</li>
	<li>0.1% polysorbate 20 (see Table of Materials) in 1x PBS (1x PBST, pH 7.4, vol/vol): to achieve a final volume of 50 mL, add 0.05 mL of polysorbate 20 to 49.95 mL of 1x PBS. Vortex and store at 4 &#176;C for up to 1 month.</li>
	<li>1% octoxynol-9 (see Table of Materials) in 1x PBS (1x PBS-TX, pH 7.4, vol/vol): to achieve a final volume of 10 mL, add 0.1 mL of octoxynol-9 to 9.9 mL of 1x PBS. Vortex and store at 4 &#176;C for up to 1 month.</li>
	<li>2% sodium azide (wt/vol): to produce 10 mL of 2% sodium azide, add 0.2 g of sodium azide to 10 mL of 1x PBS. Mix well and store at 4 &#176;C for later use.</li>
	<li>90% glycerol in 1x PBS (vol/vol): to achieve a final volume of 10 mL, add 1 mL of 1x PBS to 9 mL of glycerol. Vortex and store at room temperature (RT) for later use.</li>
</ol>
2. Animal Ddissection and Adipose Tissue Collection
<ol>
	<li>Use 6-week-old C57BL/6J male mice. Anesthetize the mice with isoflurane (4%&#8211;5% for induction then 1%&#8211;2% for maintenance). Once anesthetized, perform a toe-pinch reflex to determine the anesthetic depth, judging by the lack of pedal reflexes. Once the mice are deeply anesthetized, dissect the mice following the protocol as previously published15.</li>
	<li>Sterilize the blunt scissors and surgical area of the chest (no need to shave the hair). Open the chest by cutting the diaphragm and ribs along the lateral surface with a size of ~2 cm using blunt scissors. Clamp the chest flag with hemostat and reflect it by putting the hemostat over the head.</li>
	<li>Make a small incision into the left ventricle (~0.5 cm) with iris scissors. Insert an olive-tipped perfusion needle through the incision site and clamp the needle tip in place with the hemostat. Attach the needle to a 50 mL syringe containing paraformaldehyde fixation solution (1% PFA, pH 7.4).</li>
	<li>Open the right atrium by section cutting with iris scissors as the perfusion outlet. Start the perfusion by gently and continuously pushing the syringe at a perfusion rate close to 10 mL/min. Stop pushing when the fluid exiting the outlet is clear of any blood. The whole process should require ~5 min. 
	CAUTION: Perform these steps under a fume hood to prevent toxicity from PFA.</li>
	<li>Dissect white and brown adipose tissues from the mice. Use scissors to break the tissue samples into small pieces of chunks approximately 5 x 5&#160;x 3 mm3. Transfer the small chunks with the sterilized tweezers into the tissue embedding cassettes with proper labeling of the tissue samples on the cassettes.</li>
	<li>Immerse the embedding cassettes containing tissue samples into the fixation solution (1% PFA in 1x PBS, pH 7.4) at 4 &#176;C for 24&#8211;48 h. 
	NOTE: The fixation solution and fixation time vary based on sizes of the tissues16,17,18.</li>
	<li>Wash the samples with fresh 1x PBS buffer three times (0.5 mL/wash). 
	CAUTION: Perform this step under a fume hood. The paraformaldehyde waste should be properly handled according to hazardous waste disposal procedures. 
	NOTE: The tissues can be stored in 1x PBS at 4 &#176;C for further processing.</li>
</ol>
3. Antibody Incubation
<ol>
	<li>Cut the fixed tissue samples into approximately 2 mm3 cubes with the sterilized scissors. For permeabilization, transfer the samples into a 1.5 mL tube containing 1 mL of 1x PBS-TX (1% octoxynol-9 in 1x PBS, see Table for Materials, pH 7.4) and gently rotate the tubes at RT for 1 h at 18 rpm.</li>
	<li>Carefully remove the 1x PBS-TX by aspiration. Wash the samples 3x by adding the 1x PBS directly into the same tubes (no need to change the tubes). During each wash, invert the tubes several times.</li>
	<li>For blocking, add 0.5 mL of blocking buffer (Table of Materials) to the samples and incubate at RT for 2 h with gentle rotation.</li>
	<li>To prepare 0.4&#160;mL of primary antibody solution, dilute 2&#160;&#956;L of anti - endomucin (the marker of blood vessels) 5 (1:200) and 2&#160;&#956;L of anti &#8212; tyrosine hydroxylase (TH, the marker of nerve fibers) 5 (1:200, 1 &#181;g/mL) antibodies (Table of Materials for antibody information) into 396&#160;&#956;L of blocking buffer. Vortex and spin down to recover the volume.</li>
	<li>For the first antibody incubation, carefully remove the blocking buffer from the tissue chunks. Add 100 &#956;L of primary antibody solution prepared in step 3.4 into tubes and incubate at 4 &#176;C overnight. 
	NOTE: The antibody dilution and incubation time may vary among different antibodies. It is recommended to add 0.02% sodium azide to the antibody solution to prevent microbial growth. To prepare 0.4&#160;mL of primary antibody solution with sodium azide, add 4&#160;&#956;L of antibodies (1 &#956;L of each) and 4&#160;&#956;L of 2% sodium azide into 392&#160;&#956;L of blocking buffer.</li>
	<li>Carefully collect the primary antibody solution for reuse if desired. Wash the samples with 1x PBST (pH 7.4, 100 &#181;L/wash) three times (30 min each) with gentle rotation at 18 rpm.</li>
	<li>For the preparation of secondary antibody solution, dilute 2&#160;&#956;L of fluorophore (wave length 495 nm) conjugated anti-goat IgG (1:200) and 2&#160;&#956;L of fluorophore (wave length 650 nm) conjugated anti-rabbit IgG (1:200) (see Table of Materials) into 396&#160;&#956;L of blocking buffer. Vortex and spin briefly to collect all the liquid. 
	NOTE: Protect the samples from light during the following procedures.</li>
	<li>For the second antibody incubation, remove the last wash buffer from samples. Add 100 &#956;L of secondary antibody solution into tubes and incubate at RT for 2 h with gentle rotation at 18 rpm.</li>
	<li>Carefully remove the secondary antibody solution. Wash the samples 3x with 1x PBST (pH 7.4, 30 min each) with gentle rotation at 18 rpm. 
	NOTE: Do not reuse this secondary antibody solution, as it may be contaminated with the trace amounts of primary antibodies brought by the tissues.</li>
	<li>For optical clearance, immerse the samples in 1 mL of 90% glycerol and keep the samples at 4 &#176;C in darkness until they become transparent. 
	NOTE: The immersion duration depends on the tissue type and size. Typically, a 2 mm3 cube of white adipose tissue needs overnight incubation, while the same size brown adipose tissue sample needs longer.</li>
	<li>Adhere a silicone isolator to the slide to create a well for volume imaging. Alternatively, attach several layers of transparent tape to the slides and cut a square out of the middle to create a well (Figure 2). 
	NOTE: Adjust the well depth to fit the sample size. In this protocol, a 1 mm well depth is used.</li>
	<li>Carefully transfer the samples into the well and fill it with the mounting medium (see Table of Materials). Lay a cover slip on the surface and seal the corners of the cover slip with high viscosity medium (see Table of Materials). Let the mounting medium cure for 24 h at RT in darkness. 
	NOTE: Ensure that no space is left between the sample and cover slip. Avoid bubbles under the cover slip. Avoid placing excessive pressure on the samples.</li>
</ol>
4. Image Acquisition
<ol>
	<li>Acquire Z-stack images (refer to steps 4.2&#8211;4.7) with the 20x objective of a confocal microscope/software.</li>
	<li>Start the system. Click on the &#60;Configuration&#62; tab and activate the argon laser and HeNe 633 laser. Click on the &#60;Acquire&#62; tab. In the &#60;Visible&#62; laser lines panel, move the corresponding intensity sliders up to choose the laser lines of 488 nm and 633 nm. Initially, the intensities can be set to 20%&#8211;30%. Select the 488/561/633 triple dichoric mirror. Activate the PMTs by clicking on the &#60;Active&#62; checkboxes, choosing PMT1 for the emission excited by the 488 nm laser and PMT3 for that of the 633 nm laser. Set the wavelength range to 500&#8211;550 nm for PMT1 and 650-750nm for PMT3. Select the psudocolor to be used for image display by double clicking the colored rectangle beside each PMT and choosing a color from the pop-up menu.</li>
	<li>Click on &#60;Live&#62; to check the image of samples. Use the z-Position knob to select a plane of focus within the XY region of interest. Adjust the brightness of the images by tuning the laser intensity, smart gain and offset. For each fluorescence channel, perform this adjustment under the &#60;QLUT&#62; (Quick Look Up Table) display mode. Click on the &#60;QLUT&#62; button to enter the QLUT mode, in which saturated pixels are displayed in blue to aid the setting of appropriate brightness level. Click twice on the &#60;QLUT&#62; button to go back to the pseudocolor display mode. 
	NOTE: The numerical values of the setting may vary among each experiment.</li>
	<li>While scanning, turn the Z-Position knob counterclockwise to move the plan of focus to one end of the volume of interest and then click on the &#60;End&#62; arrowhead to set the scanning end position. Then turn the Z-Position knob clockwise to move the plane of focus through the specimen to the other end of the volume of interest and then click on the &#60;Begin&#62; arrowhead to set the scanning begin position. 
	NOTE: For the comparison between different samples, define the same Z-volume for different samples.</li>
	<li>Adjust the z-step size to 3 &#181;m. Change the image quality by selecting a format of 1024 x 1024, speed of 100 Hz, and line average of 2.</li>
	<li>Select &#60;Start&#62; to initiate the z-stack image acquisition.</li>
	<li>For maximum projection of the acquired image stack, click on &#60;Process&#62; tab and then &#60;Tool&#62;, select &#60;3D Projection&#62; and enter &#60;Maximum&#62; in the method list without changes to X, Y and Z. Set &#60;Threshold&#62; to 0. Click on &#60;Apply&#62;. The maximum intensity of the Z-volume will be stacked into a 2D image which will be shown&#160;(Figure 4A).</li>
</ol>
5. Analysis of Blood Vessel and Nerve Fiber Networks
<ol>
	<li>Analysis of 2D images via open source software (see Table of Materials)19
	<ol>
		<li>Open the stacked images in the software. Adjust Vessel diameter and intensity until all the vessel structures can be properly selected (Figure&#160;4B).</li>
		<li>Select Run analysis and export the data following guidance of the software.</li>
	</ol>
	</li>
	<li>Analysis of 3D images via the licensed software (see Table of Materials)
	<ol>
		<li>Open the raw data of images with the software. Adjust the threshold and other parameters until the vessel structures in each layer of the z-volume are properly selected (refer to the user guide for details in the software) (Figure 4C).</li>
		<li>Analyze the selected segments characters and export the data following the guidance of the software.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

Adipose tissue remodeling is directly linked to metabolic dysregulation during obesity development1,2. Angiogenesis and sympathetic innervation are both essential for the dynamic remodeling process2,12. Therefore, developing an applicable approach to visualize the new blood vessels as well as nerve fibers are of great importance. Previous methods have been reported for documenting angiogenesis in adipose tissue. However, some issues remain with these approaches, including low efficiency, fussy resolution, and noisy background. Meanwhile, sympathetic innervation has only been recognized recently as a critical step in adipose tissue pathology14. Even though the related findings are interesting, applied methods require advanced microscopy tools and are time-consuming. Here, we report an easy-to-follow approach modified from our previous protocol for co-staining blood vessels and sympathetic nerve fibers. Interestingly, we successfully stained the blood vessels and nerve fibers with high resolution, and the staining has qualities comparable to previously reported images18.
We previously stained the blood vessels in adipose tissues by immunohistochemistry or immunofluorescence with the &#945;-CD31 antibody, a marker of endothelial cells, on paraffin-embedded slides8,13,20. While the method allows us to distinguish blood vessel density between different groups, the microvascular vessels that were positively stained were not complete. Here, we chose a whole-mount method and performed IF staining with a &#945;-endomucin antibody, another marker for blood vessels. With this optimized approach, we were able to achieve staining with more blood vessels, especially microblood vessels. Moreover, in this method, since we avoided steps such as paraffin embedding and formalin fixation, which have the potential to impair the integrity of blood vessels, we obtained images with higher resolution and more intact vessels. The unimpaired structure of the vessels further allowed us to measure and compare their lengths, branching, and areas. Of note, we added a clearing step with glycerol, so the tissue chunks could become more transparent, and this simple but critical step significantly enhanced efficiency of the staining18,21.
Due to high levels of lipid contents in the adipose tissue, which might limit the accessibility of antibodies to the inner regions of the tissue, we cut the depots into small pieces to ensure adequate antibody binding to the target proteins. Therefore, our method successfully stains more blood vessels and nerve fibers than staining on whole tissues. However, it may lose spatial information and hence affect the integrity of the nerve fibers. To resolve this apparent issue and ensure the images are comparable among individual mice, it is suggested to collect the small chunks from the same regions in the adipose depots. If more spatial information is further needed, larger chunks should be obtained for staining. For larger samples, longer fixation times with higher doses of FPA, higher concentrations of detergents for permeabilization, and longer periods of antibody incubation may be needed.
Adipose tissue innervation has been investigated recently. Multiple advanced techniques have been applied to visualize nerve fibers14,17. These methods are either expensive or time-consuming. Here, we simply performed an IF stain with &#945;-TH (a sympathetic nerve marker) and successfully stained the sympathetic nerve fibers at a high quality. More importantly, we performed the co-staining with &#945;-endomucin and &#945;-TH to investigate how the blood vessels interplay with sympathetic nerve fibers during adipose tissue remodeling.
Of note, we compared the results from 2D and 3D analyses using different software. It was found that, while the 3D images provided more detailed structural information, the two analytic methods did not show significant difference in terms of the length and branching of the vessels. Eventually, when analyzing with the 3D method, the software itself may detect some false signals that need to be manually adjusted. Given that the 2D software is purposely designed for vessel analysis, it is thus suggested to use this method for quantitatively measuring the structure of vessels.
With this method, blood vessels and nerve fibers in different adipose tissue were co-stained, and their densities, branching, and lengths were compared19. It was found that both blood vessels and nerve fibers are much thicker in BAT compared to WAT, suggesting that BAT is a more metabolically active adipose depot. Moreover, the blood vessel and nerve densities in sWAT were higher than in the eWAT, indicating that different WATs have different remodeling profiles.
In conclusion, this method efficiently co-stains blood vessels and nerve fibers in adipose tissue5. Furthermore, it serves as a useful tool for studying the dynamic changes in adipose tissue during obesity development.

Adipose tissue has key metabolic and endocrine functions1. It dynamically expands or shrinks in response to different nutrient stresses2. The active tissue remodeling process consists of multiple physiological paths/steps including angiogenesis, fibrosis, and shaping of local inflammatory microenvironments2,3,4. Some physical stimuli, such as cold exposure and exercise, may trigger sympathetic activation, which ultimately leads to new blood vessel formation and sympathetic innervation in adipose tissues5,6. These remodeling processes are linked tightly to systemic metabolic outcomes including insulin sensitivity, the hallmark of type 2 diabetes2. Thus, visualization of these pathological changes is very important to understanding the healthy status of whole adipose tissues.
Angiogenesis is the process of new blood vessel formation. Since blood vessels provide oxygen, nutrients, hormones, and growth factors to tissue, angiogenesis has been considered a key step in adipose tissue remodeling, which has been documented with different techniques6,7,8,9,10,11,12,13. However, there remain questions about the resolution of the images, efficiency of immunostaining, and methods for quantification of vessel density. Compared to new blood vessel formation, innervation in adipose tissue has been underestimated for a long time. Recently, Zeng et al.14 used advanced intravital two-photon microscopy and demonstrated that adipocytes are surrounded by layers of nerve fibers14. Since then, researchers have started to appreciate the pivotal role of sympathetic innervation in regulation of adipose tissue physiology. Thus, developing an easy and practical approach to document adipose nerve innervation is important.
Here, we report an optimized method for the co-staining of blood vessels and nerve fibers based on our previous protocols. With this method, we can achieve clear images of blood vessels and nerve fibers without noisy background. Moreover, we obtain a resolution that is high enough for performing quantitative measurement of densities with open source software. By using this new approach, we can successfully compare the structures and densities of blood vessels and nerve fibers in different adipose depots.

<table>
	<tbody>
		<tr>
			<td>Alexa Fluor 488 AffiniPure Bovine Anti-Goat IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>805-545-180</td>
			<td>Lot: 116969</td>
		</tr>
		<tr>
			<td>Alexa Fluor 647 AffiniPure Donkey Anti-Rabbit IgG (H+L)</td>
			<td>Jackson ImmunoResearch</td>
			<td>711-605-152</td>
			<td>Lot: 121944</td>
		</tr>
		<tr>
			<td>Amira 6.0</td>
			<td>Thermo Fisher Scientific</td>
			<td></td>
			<td>Licensed software</td>
		</tr>
		<tr>
			<td>Angio tool</td>
			<td>National Institutes of Health</td>
			<td></td>
			<td>Open source software 
			https://ccrod.cancer.gov/confluence/display/ROB2/Home</td>
		</tr>
		<tr>
			<td>Anti-mouse endomucin antibody</td>
			<td>R&#38;D research system</td>
			<td>AF4666</td>
			<td>Lot: CAAS0115101</td>
		</tr>
		<tr>
			<td>Anti-tyrosine hydroxylase antibody</td>
			<td>Pel Freez Biologicals</td>
			<td>P40101-150</td>
			<td>Lot: aj01215y</td>
		</tr>
		<tr>
			<td>Cover glasses high performance, D=0.17mm</td>
			<td>Zeiss</td>
			<td>474030-9020-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Cytoseal 280</td>
			<td>Thermo Fisher Scientific</td>
			<td>8311-4</td>
			<td>High-viscosity medium</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Fisher</td>
			<td>G33-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde,16%</td>
			<td>TED PELLA</td>
			<td>170215</td>
			<td></td>
		</tr>
		<tr>
			<td>Press-to-Seal Silicone Isolator with Adhesive, eight wells, 9 mm diameter, 1.0 mm deep</td>
			<td>INVITROGEN</td>
			<td>P24744</td>
			<td>Silicone isolator</td>
		</tr>
		<tr>
			<td>ProLong Diamond Antifade Mountant</td>
			<td>Thermo Fisher Scientific</td>
			<td>P36965</td>
			<td>Mounting medium</td>
		</tr>
		<tr>
			<td>SEA BLOCK Blocking Buffer</td>
			<td>Thermo Fisher Scientific</td>
			<td>37527X3</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Sigma-Aldrich</td>
			<td>S2002-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Path IV Tissue Cassettes</td>
			<td>Thermo Fisher Scientific</td>
			<td>22-272416</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton &#935;-100</td>
			<td>Sigma-Aldrich</td>
			<td>X100</td>
			<td>Generic term: octoxynol-9</td>
		</tr>
		<tr>
			<td>Tube rotator and rotisseries</td>
			<td>VWR</td>
			<td>10136-084</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Sigma-Aldrich</td>
			<td>P1379</td>
			<td>Generic term: Polysorbate 20</td>
		</tr>
	</tbody>
</table>

co-staining blood vessels and nerve fibers in adipose tissue

Recent studies have highlighted the critical role of angiogenesis and sympathetic innervation in adipose tissue remodeling during the development of obesity. Therefore, developing an easy and efficient method to document the dynamic changes in adipose tissue is necessary. Here, we describe a modified immunofluorescent approach that efficiently co-stains blood vessels and nerve fibers in adipose tissues. Compared to traditional and recently developed methods, our approach is relatively easy to follow and more efficient in labeling the blood vessels and nerve fibers with higher densities and less background. Moreover, the higher resolution of the images further allows us to accurately measure the area of the vessels, amount of branching, and length of the fibers by open source software. As a demonstration using our method, we show that brown adipose tissue (BAT) contains higher amounts of blood vessels and nerve fibers compared to white adipose tissue (WAT). We further find that among the WATs, subcutaneous WAT (sWAT) has more blood vessels and nerve fibers compared to epididymal WAT (eWAT). Our method thus provides a useful tool for investigating adipose tissue remodeling.

The distal region of the epididymal white adipose tissue (eWAT), medial region of the dorsolumbar subcutaneous white adipose tissue (sWAT), and medial region of the interscapular brown adipose tissue (BAT) were collected. The locations for collecting these tissues are indicated in Figure 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59266/59266fig1.jpg" /> 
Figure 1: Anatomy of subcutaneous white adipose tissue (sWAT), epididymal white adipose tissue (eWAT), and brown adipose tissue (BAT) indicates regions used for collecting the samples. (A) The regions outlined by white dashed lines represent sWAT, and the white asterisk highlighted location is the site for collecting sWAT. Regions outlined by black dashed lines are eWAT, and the black asterisk highlighted location is the site for collecting eWAT. (B) The regions outlined by black dashed lines are BAT, and the tissue collection region is highlighted by the asterisk. <a href="https://www.jove.com/files/ftp_upload/59266/59266fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
After whole-mount staining, the tissue chunks were mounted in a well of 1 mm depth (Figure 2)
and imaged with the confocal microscope. We first tested the effect of the clearing step with 90% glycerol incubation on quality of the images. We found that more blood vessels were positively stained with &#945;-endomucin antibody in the glycerol-incubated sWAT, suggesting that the clearing step is critical for complete staining of the blood vessels (Figure 3, compare panels A and B).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59266/59266fig2.jpg" /> 
Figure 2: Diagrams of the wells for volume imaging. (A) Diagram of a slide with a silicone isolator. (B) Diagram of a well with multiple layers of tape made by our lab. <a href="https://www.jove.com/files/ftp_upload/59266/59266fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59266/59266fig3.jpg" /> 
Figure 3: Comparison of IF images of blood vessels acquiredwith or without optical clearing step with 90% glycerol. (A) Whole-mount immunofluorescence (IF) staining with anti-endomucin antibody (green) in sWAT. The sample was not subjected to the optical clearing step (step 3.10). (B) Whole-mount IF staining with anti-endomucin antibody in sWAT. The sample was subjected to the optical clearing step (step 3.10) before the mounting steps (steps 3.11 and 3.12). <a href="https://www.jove.com/files/ftp_upload/59266/59266fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
However, the clearing step with glycerol did not affect the integrity or shape of the blood vessels (Figure 3). Given these beneficial effects, in the following experiments, the clearing step was performed.
We further compared the results of 2D and 3D analyses using different software (see Table of Materials). We found that, while the 3D images apparently provided more detailed structural information, the two analytic methods eventually did not show significant differences in terms of the length and branch number of the vessels (Figure 4).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59266/59266fig4.jpg" /> 
Figure 4: Comparison of 2D and 3D analyses on vessel structure. (A) The effect of maximum projection on the images. (B) The analytic results by the 2D software (see Table of Materials). Particularly, the red lines indicate selected skeleton, while the blue dots indicate branding points. (C) The analytic results by the 3D software (see Table of Materials). The gray area is the selected region for analysis. The green line indicates the measured segments and green dots indicate the measured nodes. (D) The comparison of vessel length, segment numbers, branching node numbers, and terminal node numbers analyzed by 2D or 3D methods. <a href="https://www.jove.com/files/ftp_upload/59266/59266fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Previous publications have demonstrated that different adipose depots possess heterogeneous properties1,18. Here we sought to determine whether blood vessels and nerve fibers exhibit different patterns among the depots using our newly developed method. To achieve this, we performed co-IF staining with &#945;-endomucin (for blood vessels) and &#945;-TH (for nerve fibers) antibodies in adipose tissue. Interestingly, results showed that there were significantly more blood vessels and nerve fibers in BAT compared to WATs (Figure 5, compare bottom lanes to top and middle lanes)
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59266/59266fig5.jpg" /> 
Figure 5: Comparison of blood vessels and nerve fibers acquired from different adipose depots. Whole-mount IF staining with anti-endomucin (green) and anti-tyrosine hydroxylase (TH) (red) antibodies in eWAT (A, B, merged in C),sWAT (D, E, merged in F) and BAT (G, H, merged in I). <a href="https://www.jove.com/files/ftp_upload/59266/59266fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Among the WATs, the sWAT exhibited higher blood vessel density than the eWAT (Figure 5, compare the middle to top lane). Of note, while the nerve fibers expanded in parallel with the blood vessels, they did not show significant co-localization (Figure 5, merged lanes). We further quantitatively measured the vessel area, number of junctions, and tube length with the 2D method and found similar results as described above (Figure 6).
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59266/59266fig6.jpg" /> 
Figure 6: Quantitative analysis of vascular and nerve fiber networks. (A) The percentage of blood vessel area in the samples of eWAT, sWAT, and BAT. (B) The number of junctions of the vascular networks in eWAT, sWAT, and BAT. (C) The total vessel length of vascular networks in eWAT, sWAT, and BAT. (D) The percentage of neve fiber area in the samples of eWAT, sWAT and BAT. (E) The number of junctions of the nerve fibers in eWAT, sWAT, and BAT. (F) The total length of nerve fibers in eWAT, sWAT, and BAT. The analyses were performed with 2D software. The results were achieved from the images presented in Figure 5. <a href="https://www.jove.com/files/ftp_upload/59266/59266fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In summary, this approach successfully co-stained blood vessels and nerve fibers in different adipose tissues.

Watch this Scientific Journal Video about Co-staining Blood Vessels and Nerve Fibers in Adipose Tissue at JoVE.com

Co-staining Blood Vessels and Nerve Fibers in Adipose Tissue

New blood vessel formation and sympathetic innervation play pivotal roles in adipose tissue remodeling. However, there remain technical issues in visualizing and quantitatively measuring adipose tissue. Here we present a protocol to successfully label and quantitatively compare the densities of blood vessels and nerve fibers in different adipose tissues.

Recent studies have highlighted the critical role of angiogenesis and sympathetic innervation in adipose tissue remodeling during the development of ...

co-staining-blood-vessels-and-nerve-fibers-in-adipose-tissue

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

9.0K Views.  University of Texas Health Science Center at Houston. New blood vessel formation and sympathetic innervation play pivotal roles in adipose tissue remodeling. However, there remain technical issues in visualizing and quantitatively measuring adipose tissue. Here we present a protocol to successfully label and quantitatively compare the densities of blood vessels and nerve fibers in different adipose tissues.

Video: Co-staining Blood Vessels and Nerve Fibers in Adipose Tissue

Depletion of Specific Cell Populations by Complement Depletion

The purification of immune cell populations is often required in order to study their unique functions. In particular, molecular approaches such as real-time PCR and microarray analysis require the isolation of cell populations with high purity. Commonly used purification strategies include fluorescent activated cell sorting (FACS), magnetic bead separation and complement depletion. Of the three strategies, complement depletion offers the advantages of being fast, inexpensive, gentle on the cells and a high cell yield. The complement system is composed of a large number of plasma proteins that when activated initiate a proteolytic cascade culminating in the formation of a membrane-attack complex that forms a pore on a cell surface resulting in cell death1. The classical pathway is activated by IgM and IgG antibodies and was first described as a mechanism for killing bacteria. With the generation of monoclonal antibodies (mAb), the complement cascade can be used to lyse any cell population in an antigen-specific manner. Depletion of cells by the complement cascade is achieved by the addition of complement fixing antigen-specific antibodies and rabbit complement to the starting cell population. The cells are incubated for one hour at 37&deg;C and the lysed cells are subsequently removed by two rounds of washing. MAb with a high efficiency for complement fixation typically deplete 95-100% of the targeted cell population. Depending on the purification strategy for the targeted cell population, complement depletion can be used for cell purification or for the enrichment of cell populations that then can be further purified by a subsequent method.

To effectively study the function of immune cell populations their purification is often required. Complement depletion is a fast and inexpensive technique for the isolation of immune cell populations with high purity.

The purification of immune cell populations is often required in order to study their unique functions.  In particular, molecular approaches such as ...

Isolation of Mouse Peritoneal Cavity Cells

The peritoneal cavity is a membrane-bound and fluid-filled abdominal cavity of mammals, which contains the liver, spleen, most of the gastro-intestinal tract and other viscera. It harbors a number of immune cells including macrophages, B cells and T cells. The presence of a high number of na&iuml;ve macrophages in the peritoneal cavity makes it a preferred site for the collection of na&iuml;ve tissue resident macrophages (1). The peritoneal cavity is also important to the study of B cells because of the presence of a unique peritoneal cavity-resident B cell subset known as B1 cells in addition to conventional B2 cells. B1 cells are subdivided into B1a and B1b cells, which can be distinguished by the surface expression of CD11b and CD5. B1 cells are an important source of natural IgM providing early protection from a variety of pathogens (2-4). These cells are autoreactive in nature (5), but how they are controlled to prevent autoimmunity is still not understood completely. On the contrary, CD5+ B1a cells possess some regulatory properties by virtue of their IL-10 producing capacity (6). Therefore, peritoneal cavity B1 cells are an interesting cell population to study because of their diverse function and many unaddressed questions associated with their development and regulation. The isolation of peritoneal cavity resident immune cells is tricky because of the lack of a defined structure inside the peritoneal cavity. Our protocol will describe a procedure for obtaining viable immune cells from the peritoneal cavity of mice, which then can be used for phenotypic analysis by flow cytometry and for different biochemical and immunological assays.

The peritoneal cavity in mammals contains different immune cell populations crucial for innate immune responses. An efficient isolation method is required for biochemical and functional analyses of these cells. Here we provide a comprehensive method for the isolation of peritoneal cavity cells in the mouse.

The peritoneal cavity is a membrane-bound and fluid-filled abdominal cavity of mammals, which contains the liver, spleen, most of the gastro-intestinal ...

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting (FACS)

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known phenotype.  Other bulk methods of purification include panning, complement depletion and magnetic bead separation.  However, FACS has several advantages over other available methods.  FACS is the preferred method when very high purity of the desired population is required, when the target cell population expresses a very low level of the identifying marker or when cell populations require separation based on differential marker density.  In addition, FACS is the only available purification technique to isolate cells based on internal staining or intracellular protein expression, such as a genetically modified fluorescent protein marker.  FACS allows the purification of individual cells based on size, granularity and fluorescence.  In order to purify cells of interest, they are first stained with fluorescently-tagged monoclonal antibodies (mAb), which recognize specific surface markers on the desired cell population (1).  Negative selection of unstained cells is also possible.  FACS purification requires a flow cytometer with sorting capacity and the appropriate software.  For FACS, cells in suspension are passed as a stream in droplets with each containing a single cell in front of a laser.  The fluorescence detection system detects cells of interest based on predetermined fluorescent parameters of the cells.  The instrument applies a charge to the droplet containing a cell of interest and an electrostatic deflection system facilitates collection of the charged droplets into appropriate collection tubes (2).  The success of staining and thereby sorting depends largely on the selection of the identifying markers and the choice of mAb.  Sorting parameters can be adjusted depending on the requirement of purity and yield.  Although FACS requires specialized equipment and personnel training, it is the method of choice for isolation of highly purified cell populations.

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS

For many scientific studies requiring a biological and chemical analysis of cell populations the cells must be in a high state of purity. Fluorescence activated cell sorting (FACS) is a superior method in which to obtain pure cell populations.

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known ...

Protocol for Recombinant RBD-based SARS Vaccines: Protein Preparation, Animal Vaccination and Neutralization Detection

Based on their safety profile and ability to induce potent immune responses against infections, subunit vaccines have been used as candidates for a wide variety of pathogens 1-3. Since the mammalian cell system is capable of post-translational modification, thus forming properly folded and glycosylated proteins, recombinant proteins expressed in mammalian cells have shown the greatest potential to maintain high antigenicity and immunogenicity 4-6.
Although no new cases of SARS have been reported since 2004, future outbreaks are a constant threat; therefore, the development of vaccines against SARS-CoV is a prudent preventive step and should be carried out. The RBD of SARS-CoV S protein plays important roles in receptor binding and induction of specific neutralizing antibodies against virus infection 7-9. Therefore, in this protocol, we describe novel methods for developing a RBD-based subunit vaccine against SARS. Briefly, the recombinant RBD protein (rRBD) was expressed in culture supernatant of mammalian 293T cells to obtain a correctly folded protein with proper conformation and high immunogenicity 6. The transfection of the recombinant plasmid encoding RBD to the cells was then performed using a calcium phosphate transfection method 6,10 with some modifications. Compared with the lipid transfection method 11,12, this modified calcium phosphate transfection method is cheaper, easier to handle, and has the potential to reach high efficacy once a transfection complex with suitable size and shape is formed 13,14. Finally, a SARS pseudovirus neutralization assay was introduced in the protocol and used to detect the neutralizing activity of sera of mice vaccinated with rRBD protein. This assay is relatively safe, does not involve an infectious SARS-CoV, and can be performed without the requirement of a biosafety-3 laboratory 15.
The protocol described here can also be used to design and study recombinant subunit vaccines against other viruses with class I fusion proteins, for example, HIV, respiratory syncytial virus (RSV), Ebola virus, influenza virus, as well as Nipah and Handra viruses. In addition, the methods for generating a pseudovirus and subsequently establishing a pseudovirus neutralization assay can be applied to all these viruses.

This protocol describes a general procedure for studying recombinant receptor-binding domain (RBD)-based subunit vaccines against SARS. It includes methods for transfection and expression of RBD protein in 293T cells, immunization of mice with RBD and detection of neutralization activity of mouse sera using an established SARS pseudovirus neutralization assay.

Based on their safety profile and ability to induce potent immune responses against infections, subunit vaccines have been used as candidates for a wide ...

Introducing Shear Stress in the Study of Bacterial Adhesion

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the initial and determining step of the pathogenesis. Although experimentally adhesion is mostly studied in static conditions adhesion actually takes place in the presence of flowing liquid. First encounters between bacteria and their host often occur at the mucosal level, mouth, lung, gut, eye, etc. where mucus flows along the surface of epithelial cells. Later in infection, pathogens occasionally access the blood circulation causing life-threatening illnesses such as septicemia, sepsis and meningitis. A defining feature of these infections is the ability of these pathogens to interact with endothelial cells in presence of circulating blood. The presence of flowing liquid, mucus or blood for instance, determines adhesion because it generates a mechanical force on the pathogen. To characterize the effect of flowing liquid one usually refers to the notion of shear stress, which is the tangential force exerted per unit area by a fluid moving near a stationary wall, expressed in dynes/cm2. Intensities of shear stress vary widely according to the different vessels type, size, organ, location etc. (0-100 dynes/cm2). Circulation in capillaries can reach very low shear stress values and even temporarily stop during periods ranging between a few seconds to several minutes 1. On the other end of the spectrum shear stress in arterioles can reach 100 dynes/cm2 2. The impact of shear stress on different biological processes has been clearly demonstrated as for instance during the interaction of leukocytes with the endothelium 3. To take into account this mechanical parameter in the process of bacterial adhesion we took advantage of an experimental procedure based on the use of a disposable flow chamber 4. Host cells are grown in the flow chamber and fluorescent bacteria are introduced in the flow controlled by a syringe pump. We initially focused our investigations on the bacterial pathogen Neisseria meningitidis, a Gram-negative bacterium responsible for septicemia and meningitis. The procedure described here allowed us to study the impact of shear stress on the ability of the bacteria to: adhere to cells 1, to proliferate on the cell surface 5and to detach to colonize new sites 6 (Figure 1). Complementary technical information can be found in reference 7. Shear stress values presented here were chosen based on our previous experience1 and to represent values found in the literature. The protocol should be applicable to a wide range of pathogens with specific adjustments depending on the objectives of the study.

During the infection process, a key step is the adhesion of pathogens with host cells. In most instances this adhesion step occurs in the presence of mechanical stress generated by flowing liquid. We describe a technique that introduces shear stress as an important parameter in the study of bacterial adhesion.

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the ...

Antigens Protected Functional Red Blood Cells By The Membrane Grafting Of Compact Hyperbranched Polyglycerols

Red blood cell (RBC) transfusion is vital for the treatment of a number of acute and chronic medical problems such as thalassemia major and sickle cell anemia 1-3. Due to the presence of multitude of antigens on the RBC surface (~308 known antigens 4), patients in the chronic blood transfusion therapy develop alloantibodies due to the miss match of minor antigens on transfused RBCs 4, 5. Grafting of hydrophilic polymers such as polyethylene glycol (PEG) and hyperbranched polyglycerol (HPG) forms an exclusion layer on RBC membrane that prevents the interaction of antibodies with surface antigens without affecting the passage of small molecules such as oxygen ,glucose, and ions3. At present no method is available for the generation of universal red blood donor cells in part because of the daunting challenge presented by the presence of large number of antigens (protein and carbohydrate based) on the RBC surface and the development of such methods will significantly improve transfusion safety, and dramatically improve the availability and use of RBCs. In this report, the experiments that are used to develop antigen protected functional RBCs by the membrane grafting of HPG and their characterization are presented. HPGs are highly biocompatible compact polymers 6, 7, and are expected to be located within the cell glycocalyx that surrounds the lipid membrane 8, 9 and mask RBC surface antigens10, 11.

The cell membrane modification of red blood cells (RBCs) with hyperbranched polyglycerol (HPG) is presented. Modified RBCs were characterized by aqueous two phase partitioning, osmotic fragility and complement mediated lysis. The camouflage of surface proteins and antigens was evaluated using the flow cytometry and Micro Typing System (MTS) blood phenotyping cards.

Red blood cell (RBC) transfusion is vital for the treatment of a number of acute and chronic medical problems such as thalassemia major and sickle cell ...

Isolation of Adipose Tissue Immune Cells

The discovery of increased macrophage infiltration in the adipose tissue (AT) of obese rodents and humans has led to an intensification of interest in immune cell contribution to local and systemic insulin resistance. Isolation and quantification of different immune cell populations in lean and obese AT is now a commonly utilized technique in immunometabolism laboratories; yet extreme care must be taken both in stromal vascular cell isolation and in the flow cytometry analysis so that the data obtained is reliable and interpretable. In this video we demonstrate how to mince, digest, and isolate the immune cell-enriched stromal vascular fraction. Subsequently, we show how to antibody label macrophages and T lymphocytes and how to properly gate on them in flow cytometry experiments. Representative flow cytometry plots from low fat-fed lean and high fat-fed obese mice are provided. A critical element of this analysis is the use of antibodies that do not fluoresce in channels where AT macrophages are naturally autofluorescent, as well as the use of proper compensation controls.

Adipose tissue (AT) is a site of intense immune cell activation and interaction. Almost all cells of the immune system are present in AT and their ratios are altered by obesity. Proper isolation, quantification, and characterization of AT immune cell populations are critical for understanding their role in immunometabolic disease.

The discovery of increased macrophage infiltration in  the adipose tissue (AT) of obese rodents and humans has led to an  intensification of interest in ...

Culture of Macrophage Colony-stimulating Factor Differentiated Human Monocyte-derived Macrophages

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. For initiation of experiments, fresh or frozen monocytes are cultured in flasks for 1 week with M-CSF to induce their differentiation into macrophages. Then, the macrophages can be harvested and seeded into culture wells at required cell densities for carrying out experiments. The use of defined numbers of macrophages rather than defined numbers of monocytes to initiate macrophage cultures for experiments yields macrophage cultures in which the desired cell density can be more consistently attained. Use of cryopreserved monocytes reduces dependency on donor availability and produces more homogeneous macrophage cultures.

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. The protocol utilizes cryopreservation of monocytes coupled with their bulk differentiation into macrophages. Then harvested macrophages can then be seeded into culture wells at required cell densities for carrying out experiments.

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. For initiation ...

Flow Cytometric Analysis of Natural Killer Cell Lytic Activity in Human Whole Blood

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and reproducibility of this assay can be considered unstable, either because of user&#39;s errors or because of the sensitivity of NK cells to experimental manipulation. To eliminate these issues, a workflow that reduces them to a minimum was established and is presented here. To illustrate, we obtained blood samples, at various time points, from runners (n = 4) that were submitted to an intense bout of exercise. First, NK cells are simultaneously identified and isolated through CD56 tagging and magnetic-based sorting, directly from whole blood and from as little as one milliliter. The sorted NK cells are removed of any reagent or capping antibodies. They can be counted in order to establish an accurate NK cell count per milliliter of blood. Secondly, the sorted NK cells (effectors cells or E) can be mixed with 3,3&#39;-Diotadecyloxacarbocyanine Perchlorate (DiO) tagged K562 cells (target cells or T) at an assay-optimal 1:5 T:E ratio, and analyzed using an imaging flow-cytometer that allows for the visualization of each event and the elimination of any false positive or false negatives (such as doublets or effector cells). This workflow can be completed in about 4 h, and allows for very stable results even when working with human samples. When available, research teams can test several experimental interventions in human subjects, and compare measurements across several time points without compromising the data&#39;s integrity.

This work describes an advanced workflow for the accurate and fast determination of NK (Natural Killer) cell count and NK cell cytotoxicity in human blood samples.

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and ...

Use of a Monocyte Monolayer Assay to Evaluate Fc&#947; Receptor-mediated Phagocytosis

Although originally developed for predicting transfusion outcomes of serologically incompatible blood, the monocyte monolayer assay (MMA) is a highly versatile in vitro assay that can be modified to examine different aspects of antibody and Fc&#947; receptor (Fc&#947;R)-mediated phagocytosis in both research and clinical settings. The assay utilizes adherent monocytes from peripheral blood mononuclear cells isolated from mammalian whole blood. MMA has been described for use in both human and murine investigations. These monocytes express Fc&#947;Rs (e.g., Fc&#947;RI, Fc&#947;RIIA, Fc&#947;RIIB, and Fc&#947;RIIIA) that are involved in immune responses. The MMA exploits the mechanism of Fc&#947;R-mediated interactions, phagocytosis in particular, where antibody-sensitized red blood cells (RBCs) adhere to and/or activate Fc&#947;Rs and are subsequently phagocytosed by the monocytes. In vivo, primarily tissue macrophages found in the spleen and liver carry out Fc&#947;R-mediated phagocytosis of antibody-opsonized RBCs, causing extravascular hemolysis. By evaluating the level of phagocytosis using the MMA, different aspects of the in vivo Fc&#947;R-mediated process can be investigated. Some applications of the MMA include predicting the clinical relevance of allo- or autoantibodies in a transfusion setting, assessing candidate drugs that promote or inhibit phagocytosis, and combining the assay with fluorescent microscopy or traditional Western immunoblotting to investigate the downstream signaling effects of Fc&#947;R-engaging drugs or antibodies. Some limitations include the laboriousness of this technique, which takes a full day from start to finish, and the requirement of research ethics approval in order to work with mammalian blood. However, with diligence and adequate training, the MMA results can be obtained within a 24-h turnover time.

The monocyte monolayer assay (MMA) is an in vitro assay that utilizes isolated primary monocytes obtained from mammalian peripheral whole blood to evaluate Fc&#947; receptor (Fc&#947;R)-mediated phagocytosis.

Although originally developed for predicting transfusion outcomes of serologically incompatible blood, the monocyte monolayer assay (MMA) is a highly ...