This work was supported by the NIH (R01 AI43458 to H.L.W.).

The protocol was approved by the Standing Committee on Animals at Harvard Medical School and Brigham and Women&#8217;s Hospital, protocol 2016N000230.
1. Mice used for the experiment
<ol>
	<li>Use 8-week old male and female mice on the B6 background for administering the antibody mix.</li>
	<li>Use CX3CR1GFP/WTCCR2RFP/WT mice to determine whether ex vivo whole LN imaging can also be applied to reporter mice without administering antibody mix as well as to investigate the presence of mononuclear cells, including antigen presenting cells and phagocytes, and their distribution in the LN. 
	NOTE: CX3CR1GFP/WTCCR2RFP/WT reporter mice have green fluorescent protein (GFP) and red fluorescent protein (RFP) inserted under the control of CX3CR1 and CCR2 promoters, respectively. Reporter mice can be used with or without the injection of the antibody mix. Please see reference4 for antibody mix injection in a reporter mouse. Proceed to surgery if no antibody will be injected.</li>
</ol>
2. Antibody mix preparation and injection
NOTE: Perform these steps on mice described in step 1.1.
<ol>
	<li>Dilute 1:10 of brilliant violet (BV) 421 anti-CD4 (GK1.5; 0.2 mg/mL), 1:10 of brilliant blue (BB) 515 anti-CD19 (1D3; 0.2 mg/mL) and 1:20 of phycoerythrin (PE) anti-F4/80 (T45-2342; 0.2 mg/mL) in PBS with the appropriate final volume to inject into the inner thigh (to image inguinal) or into the paw pad (to image popliteal) lymph. 
	NOTE: Use as isotypes: BV421 Mouse IgG2b, k Isotype Control (R35-38; 0.2 mg/mL); PE Rat IgG2a, &#954; Isotype Control (R35-95; 0.2 mg/mL); BB515 Rat IgG2a, &#954; Isotype Control (R35-95; 0.2 mg/mL). If changing the staining antibody, use correct isotype.</li>
	<li>To image inguinal LN, inject 100 &#181;L of the antibody mix subcutaneously into the inner thigh (Figure 1A). Alternatively, inject 50 &#181;L subcutaneously of antibody mix into the paw pad to image popliteal LN (Figure 2A). Use delicate 1 mL Insulin syringes, (Insulin U-100) with the needle of size 0.30 mm &#215; 13 mm (30 gauge &#215; &#189; inch). 
	NOTE: Ensure that the injection for inguinal LN staining is subcutaneous and not intramuscular (i.m.), as antibody mix will not be properly drained if i.m. administration is performed.</li>
	<li>Do not anesthetize animals before antibody mix injection.</li>
	<li>Wait for a minimum of 3 h (inguinal dLN) and 12 h (popliteal dLN) post injection to remove the organs.</li>
	<li>If LN cell labeling is not completely observed using large polymer fluorescent dyes such as brilliant violet or brilliant blue, use smaller fluorophores, including fluorescein isothiocyanate (FITC), PE and allophycocyanin (APC) as an alternative.</li>
</ol>
3. Surgery procedure to remove the inguinal draining lymph node
<ol>
	<li>Euthanize mice using CO2 asphyxiation followed by cervical dislocation.</li>
	<li>Immobilize mice on the acrylic stage with adhesive tape and apply mineral oil with a cotton swab to the abdominal skin to prevent fur deposition around the incision (Figure 1B). Fur removal is not necessary.</li>
	<li>Perform a midline incision using microsurgery curved scissors (11.5 cm) and microsurgery curved forceps (12.5 cm) in the abdomen from the pubis to the xiphoid process (Figure 1C).</li>
	<li>Dissociate the abdominal musculature from the skin (Figure 1D).</li>
	<li>Make horizontal skin incisions at the top and bottom of the vertical incision line to create skin flaps on side of interest (according to the side of the antibody mix injection) and flap the skin to visualize the lymph node (Figure 1E).</li>
	<li>Tape the skin-flap on the acrylic plate (Figure 1F).</li>
	<li>Remove the inguinal draining lymph node using microsurgery curved forceps (Figure 1F). Lymph node will appear as a translucid, usually bilobular, sphere under the skin.</li>
</ol>
4. Surgery procedure to remove the popliteal draining lymph node
<ol>
	<li>Euthanize mice using CO2 asphyxiation followed by cervical dislocation.</li>
	<li>Immobilize mice at a prone position on acrylic stage with adhesive tape and apply mineral oil with a cotton swab in the calf and knee (Figure 2A-D).</li>
	<li>Perform a midline incision in the calf from the heel to the knee (Figure 2E,F).</li>
	<li>Dissociate the calf musculature from the skin (Figure 2F,G).</li>
	<li>Expose the popliteal fossa (Figure 2G). Popliteal lymph node will appear as a translucid sphere in the popliteal fossa.</li>
	<li>Remove the popliteal lymph node using microsurgery curved forceps (Figure 2G).</li>
	<li>Alternatively, turn the mouse over on supine position, and approach the popliteal fossa between biceps femoris and semitendinosus for popliteal LN removal by performing a midline incision in the calf from the heel to the knee followed by dissociation of the calf musculature from the skin. 
	NOTE: Popliteal fossa is a shallow depression located at the back of the knee joint. Open carefully to see the popliteal lymph node.</li>
</ol>
5. Lymph node preparation
<ol>
	<li>Place the whole organ in a culture dish with glass bottom (35 mm x 10 mm) and remove the fat that surrounds the organ using microsurgery curved forceps (Figure 1G-I and Figure 2H).</li>
	<li>Centralize the organ in the middle of the dish (Figure 1J and Figure 2H).</li>
	<li>Cover the organ with a fragment of delicate task wipers and keep it soaked with room temperature saline 0.9% or Phosphate Buffer Solution (Figure 1K-M and Figure 2H). 
	NOTE: It is not necessary to wash lymph nodes after removal or to perform the lymph node extraction in the hood.</li>
</ol>
6. Ex-vivo confocal microscopy (Imaging)
<ol>
	<li>Position the Petri dish in the inverted confocal microscope slot (Figure 1N and Figure 2H).</li>
	<li>Image LNs under a confocal microscope (Figure 1O and Figure 2H).</li>
	<li>First, obtain the correct focus by using the conventional light of the confocal microscope with 4x or 10x objective. Then change from the light function to the laser mode. 
	NOTE: If the confocal microscope does not contain a 4x objective, the focus can be perfectly obtained using the 10x objective.</li>
	<li>Adjust the laser power, offset and gain using an isotype-stained, non-stained or non-fluorescent sample (Table 1) to remove autofluorescence and unspecific staining from fluorescent-labeled antibodies such as the ones used in the images showed in this manuscript (BV-421 anti-CD4, BB515 anti-CD19 and PE anti-F4/80).</li>
	<li>Adjust Z and XY positions, using the micrometric and chariot, respectively, on the area of interest in the lymph node.</li>
	<li>Acquire images under 4x, 10x and 20x objectives focusing on the LN structure and cellular distribution. Use 1024 x 1024-pixel definition. 
	NOTE: Acquire a minimum of five images in different fields per objective per animal.</li>
	<li>Analyze images using an image software to separate channels, add scale and colors of interest, and to reconstruct 3D view.</li>
</ol>

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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=Combination+of+Mass+Cytometry+and+Imaging+Analysis+Reveals+Origin,+Location,+and+Functional+Repopulation+of+Liver+Myeloid+Cells+in+Mice.">Combination of Mass Cytometry and Imaging Analysis Reveals Origin, Location, and Functional Repopulation of Liver Myeloid Cells in Mice.</a> Gastroenterology. 151, 1176-1191 (2016).</li><li>Rezende, R. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=gammadelta+T+cells+control+humoral+immune+response+by+inducing+T+follicular+helper+cell+differentiation.">gammadelta T cells control humoral immune response by inducing T follicular helper cell differentiation.</a> Nature Communications. 9, 3151 (2018).</li><li>Nakagaki, B. 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The authors have nothing to disclose.

The combination of imaging with other techniques, including molecular biology and high dimensional immunophenotyping has enhanced our ability to investigate immune cells in their native context. In fact, while other approaches may require tissue digestion and cell isolation &#8211; which can lead to loss of tissue integrity - the use of in vivo or ex vivo imaging grants a great advantage in investigating different cell subtypes in a geographical fashion3,16. It i...

Lymph nodes (LNs) are ovoid-shaped organs widely present throughout the body with the crucial function of bridging the innate and adaptive immune responses. LNs filter the lymph in order to identify foreign particles and cancerous cells to mount an immune response against them1. Antigen presenting cells (APCs), T cells and B cells work alongside to generate antigen-specific antibodies (humoral immunity) and cytotoxic lymphocytes (cellular immunity) to eliminate the foreign particles and cancerous cells2. Thus, understanding the dynamics of the immune cells present in the lymphatic system will have important implications for the vaccine development and cancer immunotherapy.
The advent of powerful microscopes - including new confocal and super resolution microscopes - has allowed an extraordinary advancement in understanding how different immune cell populations behave in their native environment3. It is now possible to image several simultaneous cell subtypes using a combination of probes with genetically modified mice that express fluorescent proteins under control of specific targets4,5. In fact, high dimensional techniques, including mass cytometry and multi-parametric flow analysis have been crucial to expanding our knowledge on different immune cell compartmentalization and functionality in the health and disease6,7. However, to prepare samples for these techniques, tissues need digestion and cells are separated from their natural milieu to be analyzed in cell suspensions. To surpass these limitations and allow a better translation in biology, the goal of the protocol proposed here is to apply a straightforward methodology to image ex vivo whole lymph nodes using stock confocal microscopes with the benefit of improved speed, tissue structure preservation, and cell viability compared to the conventional immunofluorescence staining. By using this approach, we were able to show that mice deficient for &#947;&#948; T cells, a subtype of T lymphocyte involved in host early defense against pathogens4, have compromised follicles and T cell zones as compared to wild type mice. These findings allowed us to pursue a study in which we demonstrated that &#947;&#948; T cells play a critical role in the homeostasis of lymphoid organs and humoral immune response4. Furthermore, this protocol provides a physiologic pathway for probes and antibodies to reach the lymph node, as they are administered subcutaneously and dissipate through the tissue lymphatic circulation, building on previous reports that used in situ labeling with antibodies to visualize lymphatic-associated structures8,9, germinal center dynamics10,11,12, and targets readily accessible to blood flow13,14,15.

<table>
	<tbody>
		<tr>
			<td>BV421 anti-CD4</td>
			<td>BD Horizon</td>
			<td>562891</td>
			<td>GK1.5; 0.2 mg mL-1</td>
		</tr>
		<tr>
			<td>BB515 anti-CD19</td>
			<td>BD Horizon</td>
			<td>564509</td>
			<td>1D3; 0.2 mg mL-1</td>
		</tr>
		<tr>
			<td>BB515 Rat IgG2a, &#954; Isotype Control</td>
			<td>BD Horizon</td>
			<td>564418</td>
			<td>R35-95; 0.2 mg mL-1</td>
		</tr>
		<tr>
			<td>BV421 Mouse IgG2b, K Isotype Control</td>
			<td>BD Horizon</td>
			<td>562603</td>
			<td>R35-38 0.2 mg mL-1</td>
		</tr>
		<tr>
			<td>Cellview culture dish</td>
			<td>Greiner-Bio</td>
			<td>627861</td>
			<td>35x10 mm with glass bottom</td>
		</tr>
		<tr>
			<td>Insulin syringes</td>
			<td>BD Plastipak</td>
			<td>-</td>
			<td>Insulin U-100</td>
		</tr>
		<tr>
			<td>Kimwipes</td>
			<td>Kimtech Science Brand</td>
			<td>7557</td>
			<td>size 21 x 20 cm / 100 sheets per box</td>
		</tr>
		<tr>
			<td>Microsurgery curved forceps</td>
			<td>WEP Surgical Instruments</td>
			<td>custom made</td>
			<td>12.5 cm</td>
		</tr>
		<tr>
			<td>Microsurgery curved scissors</td>
			<td>WEP Surgical Instruments</td>
			<td>custom made</td>
			<td>11.5 cm</td>
		</tr>
		<tr>
			<td>Needle</td>
			<td>BD PrecisionGlide</td>
			<td>-</td>
			<td>30 gauge &#215; &#189; inch</td>
		</tr>
		<tr>
			<td>Nikon Eclipse Te + A1R confocal head</td>
			<td>Nikon</td>
			<td>-</td>
			<td>loaded with main 4 laser lines (405, 488, 543 and 647 nm)</td>
		</tr>
		<tr>
			<td>PE anti-F4/80</td>
			<td>BD Pharmigen</td>
			<td>565410</td>
			<td>T45-2342; 0.2 mg mL-1</td>
		</tr>
		<tr>
			<td>PE Rat IgG2a, &#954; Isotype Control</td>
			<td>BD Pharmigen</td>
			<td>553930</td>
			<td>R35-95; 0.2 mg mL-1</td>
		</tr>
		<tr>
			<td>Zeiss LSM 710 confocal microscope</td>
			<td>Zeiss</td>
			<td>-</td>
			<td>loaded with main 4 laser lines (405, 488, 543 and 647 nm)</td>
		</tr>
	</tbody>
</table>

visualizing lymph node structure and cellular localization using ex-vivo confocal microscopy

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are strategically interposed in the path of the lymphatic vessels, allowing intimate contact of tissue antigens with all resident immune cells in the LN. Thus, understanding the cellular composition, distribution, location and interaction using ex vivo whole LN imaging will add to the knowledge on how the body coordinates local and systemic immune responses. This protocol shows an ex vivo imaging strategy following an in vivo administration of fluorescent-labeled antibodies that allows a very reproducible and easy-to-perform methodology by using conventional confocal microscopes and stock reagents. Through subcutaneous injection of antibodies, it is possible to label different cell populations in draining LNs without affecting tissue structures that can be potentially damaged by a conventional immunofluorescence microscopy technique.

This manuscript shows techniques to remove inguinal and popliteal lymph nodes without damaging their structure following the injection of fluorescent-labeled antibodies to stain specific cell populations in these organs (Figure 1 and Figure 2).
The powerful combination of immunolabeling of LN cells with BV421 anti-CD4 and BB515 anti-CD19 and confocal imaging analysis defined the localization of T cells (CD4+) and B cells (CD19+) in in...

Watch this Scientific Journal Video about Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy at JoVE.com

Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy

生体共焦点顕微鏡を用いてリンパ節構造と細胞局在化を可視化

This protocol describes a technique to image different cell populations in draining lymph nodes without alterations in the organ structure.

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are ...

visualizing-lymph-node-structure-cellular-localization-using-ex-vivo

Research

JoVE Journal

Biology

12.9K ビュー.  Brigham and Women's Hospital, Harvard Medical School. この再現性と容易なプロトコルを実行するには、元生体リンパ節イメージング戦略を使用して、生体内投与された蛍光標識抗体の局所的な二次リンパ器官への排出を追跡します。この手法をデモンストレーションすることで、プロトコルの実行を成功に導く手順を特定の視覚化できます。これは、誰でも実行できる非常にシンプルで簡単なテクニックです。唯一の?...

ビデオ: Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

可視化RNAの局在にアフリカツメガエル卵母細胞

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

生物学

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells (HSCs) differentiate into oligopotent progenitors, committed precursors and mature red blood cells 1. This process is regulated for a large part at the level of gene expression, whereby specific transcription factors activate lineage-specific genes while concomitantly repressing genes that are specific to other cell types 2. Studies on transcription factors regulating erythropoiesis are often performed using human and murine cell lines that represent, to some extent, erythroid cells at given stages of differentiation 3-5. However transformed cell lines can only partially mimic erythroid cells and most importantly they do not allow one to comprehensibly study the dynamic changes that occur as cells progress through many stages towards their final erythroid fate. Therefore, a current challenge remains the development of a protocol to obtain relatively homogenous populations of primary HSCs and erythroid cells at various stages of differentiation in quantities that are sufficient to perform genomics and proteomics experiments.
Here we describe an ex vivo cell culture protocol to induce erythroid differentiation from human hematopoietic stem/progenitor cells that have been isolated from either cord blood, bone marrow, or adult peripheral blood mobilized with G-CSF (leukapheresis). This culture system, initially developed by the Douay laboratory 6, uses cytokines and co-culture on mesenchymal cells to mimic the bone marrow microenvironment. Using this ex vivo differentiation protocol, we observe a strong amplification of erythroid progenitors, an induction of differentiation exclusively towards the erythroid lineage and a complete maturation to the stage of enucleated red blood cells. Thus, this system provides an opportunity to study the molecular mechanism of transcriptional regulation as hematopoietic stem cells progress along the erythroid lineage. 
Studying erythropoiesis at the transcriptional level also requires the ability to over-express or knockdown specific factors in primary erythroid cells. For this purpose, we use a lentivirus-mediated gene delivery system that allows for the efficient infection of both dividing and non-dividing cells 7. Here we show that we are able to efficiently knockdown the transcription factor TAL1 in primary human erythroid cells. In addition, GFP expression demonstrates an efficiency of lentiviral infection close to 90%. Thus, our protocol provides a highly useful system for characterization of the regulatory network of transcription factors that control erythropoiesis.

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

An ex vivo protocol to generate mature human red blood cells from hematopoietic stem/progenitors is described. Additionally we describe an efficient lentiviral-delivery method to knockdown the transcription factor TAL1 in primary erythroid cells. The efficiency of lentivirus mediated gene delivery is demonstrated using GFP expressing viruses.

レンチウイルスを介したノックダウン中生体外で</emヒト造血幹細胞の>赤血球

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells ...

A System for ex vivo Culturing of Embryonic Pancreas

The pancreas controls vital functions of our body, including the production of digestive enzymes and regulation of blood sugar levels1. Although in the past decade many studies have contributed to a solid foundation for understanding pancreatic organogenesis, important gaps persist in our knowledge of early pancreas formation2. A complete understanding of these early events will provide insight into the development of this organ, but also into incurable diseases that target the pancreas, such as diabetes or pancreatic cancer. Finally, this information will generate a blueprint for developing cell-replacement therapies in the context of diabetes.
 During embryogenesis, the pancreas originates from distinct embryonic outgrowths of the dorsal and ventral foregut endoderm at embryonic day (E) 9.5 in the mouse embryo3,4. Both outgrowths evaginate into the surrounding mesenchyme as solid epithelial buds, which undergo proliferation, branching and differentiation to generate a fully mature organ2,5,6. Recent evidences have suggested that growth and differentiation of pancreatic cell lineages, including the insulin-producing &beta;-cells, depends on proper tissue-architecture, epithelial remodeling and cell positioning within the branching pancreatic epithelium7,8. However, how branching morphogenesis occurs and is coordinated with proliferation and differentiation in the pancreas is largely unknown. This is in part due to the fact that current knowledge about these developmental processes has relied almost exclusively on analysis of fixed specimens, while morphogenetic events are highly dynamic.
 Here, we report a method for dissecting and culturing mouse embryonic pancreatic buds ex vivo on glass bottom dishes, which allow direct visualization of the developing pancreas (Figure 1). This culture system is ideally devised for confocal laser scanning microscopy and, in particular, live-cell imaging. Pancreatic explants can be prepared not only from wild-type mouse embryos, but also from genetically engineered mouse strains (e.g. transgenic or knockout), allowing real-time studies of mutant phenotypes. Moreover, this ex vivo culture system is valuable to study the effects of chemical compounds on pancreatic development, enabling to obtain quantitative data about proliferation and growth, elongation, branching, tubulogenesis and differentiation. In conclusion, the development of an ex vivo pancreatic explant culture method combined with high-resolution imaging provides a strong platform for observing morphogenetic and differentiation events as they occur within the developing mouse embryo.

A System for ex vivo Culturing of Embryonic Pancreas

Here, we describe a method for isolation, culture and manipulation of mouse embryonic pancreas. This represents an excellent ex vivo system for studying various aspects of pancreatic development, including morphogenesis, differentiation and growth. Pancreatic bud explants can be cultured for several days and used in a range of different applications, including whole-mount immunofluorescence and live imaging.

のためのシステム<em&gt; ex vivoで</em胚膵臓&gt;培養

The pancreas controls vital functions of our body,  including the production of digestive enzymes and regulation of blood sugar  levels1. Although in the ...

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

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

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

ナノFEM：写真活性化ローカライゼーション顕微鏡と電子顕微鏡を用いてタンパク質の局在

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

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

蛍光顕微鏡を用いて線虫の中の細菌の可視化

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

4D共焦点顕微鏡を用いたゼブラフィッシュにおける顔面形態形成を分析する

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

共焦点および多光子顕微鏡を用いて5蛍光タンパク質によってマーク造血幹細胞および前駆細胞の生体内でクローン追跡で

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

の剛性を測定します<em&gt; ex vivoで</em原子間力顕微鏡を使用&gt;マウス大動脈

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Cellular Redox Profiling Using High-content Microscopy

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, excessive ROS levels induce a state of oxidative stress, which is accompanied by irreversible oxidative damage to DNA, lipids and proteins. Thus, quantification of ROS provides a direct proxy for cellular health condition. Since mitochondria are among the major cellular sources and targets of ROS, joint analysis of mitochondrial function and ROS production in the same cells is crucial for better understanding the interconnection in pathophysiological conditions. Therefore, a high-content microscopy-based strategy was developed for simultaneous quantification of intracellular ROS levels, mitochondrial membrane potential (&#916;&#936;m) and mitochondrial morphology. It is based on automated widefield fluorescence microscopy and image analysis of living adherent cells, grown in multi-well plates, and stained with the cell-permeable fluorescent reporter molecules CM-H2DCFDA (ROS) and TMRM (&#916;&#936;m and mitochondrial morphology). In contrast with fluorimetry or flow-cytometry, this strategy allows quantification of subcellular parameters at the level of the individual cell with high spatiotemporal resolution, both before and after experimental stimulation. Importantly, the image-based nature of the method allows extracting morphological parameters in addition to signal intensities. The combined feature set is used for explorative and statistical multivariate data analysis to detect differences between subpopulations, cell types and/or treatments. Here, a detailed description of the assay is provided, along with an example experiment that proves its potential for unambiguous discrimination between cellular states after chemical perturbation.

This paper presents a high-content microscopy workflow for simultaneous quantification of intracellular ROS levels, as well as mitochondrial membrane potential and morphology &#8211; jointly referred to as mitochondrial morphofunction &#8211; in living adherent cells using the cell-permeant fluorescent reporter molecules 5-(and-6)-chloromethyl-2&#39;,7&#39;-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) and tetramethylrhodamine methylester (TMRM).

ハイコンテンツ顕微鏡を用いた細胞レドックスプロファイリング

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.