The study was supported by grants from the National Natural Science Foundation of China (81960226, 81760223), the Natural Science Foundation of Yunnan Province (202001AS070045, 202301AY070001-011), and the Scientific Research Foundation of Yunnan Province Department of Education (2023Y0784).

This research received approval from the Animal Experiment Ethics Committee of Kunming Medical University (kmmu20220945). All experiments adhered to established animal care guidelines. Leptomeningeal lymphatic endothelial cells (LLECs) were harvested from male C57Bl/6J mice aged 6-8 weeks and weighing between 20-25 g. These mice were procured from Kunming Medical University in Kunming, China. The entire experimental procedure was conducted under strict sterile conditions. All the centrifugation steps are performed at room temperature unless otherwise specified.
1. Preparation of reagents and instruments
NOTE: All steps involving solutions must be conducted within a class II biohazard cabinet.
<ol>
	<li>Prepare the washing buffer by mixing phosphate-buffered saline (PBS) with calcium and magnesium, 10% fetal bovine serum (FBS), and 1% penicillin-streptomycin (P/S) (see Table of Materials). Keep this buffer cold at 4 &#176;C. It is essential to prepare it fresh on the day of use and degas the buffer.</li>
	<li>Prepare the digestive enzymes by combining 10 mL of PBS (without calcium and magnesium) with 2 mL of 0.25% trypsin, 1 mL of 20 mg/mL papain, and 200 &#181;L of 1 mg/mL collagenase I (see Table of Materials). 
	NOTE: Ensure the use of aliquots of appropriate volumes to prevent repeated freeze-thaw cycles. Store these aliquots at -20 &#176;C.</li>
	<li>Prepare the culture medium by utilizing the commercially available endothelial cell growth medium kit, which includes VEGF-C (100 ng/mL) and 1% P/S (see Table of Materials).</li>
	<li>Prepare the stopping buffer by supplementing Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) with 10% FBS to stop the digestion process.</li>
	<li>Prepare sterile surgical instruments: these instruments should include scissors, serrated-tip tweezers, and fine-point tweezers.</li>
</ol>
2. Flask coating
<ol>
	<li>Coat the T25 flask with a 100 &#181;g/mL fibronectin solution (see Table of Materials) and incubate overnight at 37 &#176;C. Prior to use, remove the coating solution using a pipette.</li>
	<li>Wash the flask three times with PBS, subsequently aspirate the PBS, and allow the T25 flask to air dry.</li>
</ol>
3. Leptomeninges dissociation
NOTE: Always use pre-cooled buffers and solutions at 4 &#176;C.
<ol>
	<li>Anesthetize the mice with excess inhalation of 4% isoflurane and then sacrifice them by swift decapitation using scissors that have been previously cleaned with 70% ethanol.</li>
	<li>Carefully incise the skin along the midline, starting from the opening at the back of the skull and extending toward the frontal area.</li>
	<li>Delicately remove the skull, lifting it gently with the scissors to avoid damaging the leptomeninges, ensuring the entire brain, including the leptomeninges, is obtained.</li>
	<li>Submerge the whole brain in a washing buffer and gently flush it to remove surface blood.</li>
	<li>Transfer the brain into a sterile Petri dish without chopping, and extract the leptomeninges from the brain&#39;s surface using fine-point tweezers under a microscope.</li>
	<li>Cut the leptomeninges tissue into fragments using sterile micro-scissors.</li>
</ol>
4. Leptomeninges enzymatic digestion
<ol>
	<li>Add 10 mL of the enzyme mix (step 1.2) to the fragments and incubate at 37 &#176;C for 15 min. Gently agitate to ensure the fragments detach from the bottom of the tube. Afterward, add 10 mL of stopping buffer (step 1.4) to halt the digestion.</li>
	<li>Centrifuge at 300 x g for 5 min at 4 &#176;C and carefully remove the supernatant using a pipette.</li>
	<li>Add 10 mL of cold PBS, and pass the mixture through a 70 &#181;m strainer into a sterile 50 mL tube to filter out any clumps.</li>
</ol>
5. Cell expansion
<ol>
	<li>Plate 1 x 105 cells per cm2 into a fibronectin-coated T25 flask (step 2) with 5 mL of culture medium.</li>
	<li>Incubate the cells at 37 &#176;C with 5% CO2 for 24 h. Afterward, remove the medium to eliminate non-attached cells.</li>
	<li>Sustain the culture by replacing 50% of the medium every two days. Repeat this process two or three times.</li>
</ol>
6. Magnetic cell selection
NOTE: Work swiftly, maintain cell coldness, and utilize pre-cooled solutions to prevent non-specific cell labeling.
<ol>
	<li>Preparation of instruments and reagents: attach a magnetic separator to a magnetic separator stand (see Table of Materials). Connect a selection column to the magnetic separator, and position a 70 &#181;m cell strainer atop the selection column. Place a 50 mL tube beneath the selection column to collect the flow.</li>
	<li>Enzymatic digestion: once the cells reach 80% confluence, aspirate the medium and rinse the cells with PBS. Add 0.25% trypsin to detach adherent cells and incubate at 37 &#176;C for 5 min. Stop the digestion by adding the stopping buffer. Centrifuge the cells at 300 x g for 5 min at room temperature and remove the supernatant.</li>
	<li>Antibody incubation: resuspend 1 x 107 total cells in 100 &#181;L of PBS and add 10 &#181;L of LYVE-1 antibody (see Table of Materials). Mix thoroughly and incubate for 30 min in the dark at 4 &#176;C.
	<ol>
		<li>Spin the cells at 300 x g for 5 min and discard the supernatant. Rinse the cells by introducing 1 mL of PBS and centrifuging at 300 x g for 5 min. Completely remove the supernatant.</li>
	</ol>
	</li>
	<li>Microbeads labeling: resuspend the cells in 100 &#181;L of PBS and add 20 &#181;L of Microbeads (see Table of Materials). Mix well and incubate for 30 min in the dark at 4 &#176;C.
	<ol>
		<li>Spin the cells at 300 x g for 5 min and decant the supernatant. Subsequently, cleanse the cells with 1 mL of PBS through centrifugation at 300 x g for 5 min. Finally, completely remove the supernatant.</li>
	</ol>
	</li>
	<li>Magnetic negative exclusion: resuspend the cells in 4 mL of PBS. Pass the cells through a 70 &#181;m cell strainer to eliminate clumps. Prepare the selection column (see Table of Materials) by rinsing it with 3 mL of PBS, then apply the cell suspension into the selection column.
	<ol>
		<li>Perform washing steps with 3 mL of PBS and collect LYVE-1-negative cells into a 50 mL tube positioned below the selection column to allow them to pass through.</li>
	</ol>
	</li>
	<li>Magnetic positive selection: pipette 6 mL of PBS into the selection column. Instantly flush out the magnetically labeled cells by firmly pushing the plunger into the selection column to obtain LYVE-1-positive LLECs. 
	&#8203;NOTE: Always wait until the selection column reservoir is empty before proceeding to the next step.</li>
</ol>
7. LLECs culture
<ol>
	<li>Centrifuge the LYVE-1-positive LLECs at 300 x g for 5 min, and then carefully remove the supernatant.</li>
	<li>Plate 1 x 105 cells per cm2 into a fibronectin-coated T25 flask with 5 mL of culture medium.</li>
	<li>Maintain the culture by replacing 50% of the medium every other day. When the cells reach 80% confluence, detach the cells with 0.25% trypsin and perform cell passage. Utilize the cells for subsequent experiments after 2-3 passages.</li>
</ol>
8. Flow cytometric analysis
NOTE: The Flow cytometric analysis was conducted following the previously described procedure15.
<ol>
	<li>Add 0.25% trypsin to detach adherent cells and incubate at 37 &#176;C for 5 min. Then, add the stopping buffer to halt the digestion.</li>
	<li>Centrifuge the cells at 300 x g for 5 min and completely aspirate the supernatant. Resuspend 1 x 106 cells in 100 &#181;L of PBS.</li>
	<li>Add 1 &#181;L of LYVE-1 antibody. Mix thoroughly and incubate for 10 min in the dark at room temperature.</li>
	<li>Resuspend the cells in an appropriate amount of PBS buffer for analysis using flow cytometry and FlowJo software (see Table of Materials).</li>
</ol>
9. Immunofluorescent staining
NOTE: The immunofluorescent staining was conducted following the procedure described previously16.
<ol>
	<li>Wash the cells with PBS three times. Fix the cells with 4% paraformaldehyde (PFA) for 10 min at room temperature. Discard the PFA and wash the cells with PBS three times.</li>
	<li>Apply the block buffer for 1 h at room temperature. Remove the block buffer from the coverslips.</li>
	<li>Add the LYVE-1 antibody in the staining buffer to the cells and incubate in the dark. Wash the cells three times with PBS.</li>
	<li>Add an appropriate secondary antibody (see Table of Materials) in the staining buffer to the cells and incubate in the dark. Wash the cells three times with PBS.</li>
	<li>Counterstain with 4&#39;,6-diamidino-2-phenylindole (DAPI). The cells are now ready for immunofluorescence microscopy.</li>
</ol>

Establishing In Vitro LLEC Cultures Using Magnetic Cell Selection

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L., Ruddell, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culturing+purifies+murine+lymph+node+lymphatic+endothelium.">Culturing purifies murine lymph node lymphatic endothelium.</a> Lymphatic Research and Biology. 12 (3), 144-149 (2014).</li><li>Jones, B. E., Yong, L. 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The authors declare that they have no conflict of interest to disclose.

The existing protocol for harvesting and culturing LLECs in vitro has not been previously reported. This study introduces a reproducible, multi-procedural protocol for harvesting and culturing LLECs from mouse leptomeninges.
While this multi-procedural protocol is reproducible, there are several key considerations. For example, fibronectin-coated T25 flasks promote the adhesion of LLECs and function by eliminating non-adherent cells, thereby ensuring a more homogenous cellular populat...

The newly discovered leptomeningeal lymphatic endothelial cells (LLECs) form a meshwork of individual cells within the leptomeninges, exhibiting a distinct distribution pattern when compared to peripheral lymphatic endothelial cells1,2. The cellular functions and clinical implications associated with LLECs remain largely uncharted territory. In order to pave the way for functional research on LLECs, it is imperative to establish an in vitro model for their study. Therefore, this study has devised a comprehensive protocol for the isolation and primary culture of LLECs.
Mice are the preferred animal model due to their suitability for genetic manipulation in disease research. Previous studies have successfully isolated lymphatic endothelial cells from various mouse tissues, including lymph nodes3, mesenteric tissue4, dermal tissue5, collecting lymphatics6, and lung tissue7. These isolation procedures have primarily relied on techniques such as magnetic-activated cell sorting (MACS) and flow cytometry sorting8,9,10. Additionally, research efforts have led to the establishment of rat arachnoid cell lines and rat lymphatic capillary cell lines11,12. Despite the existence of explant culture techniques for leptomeninges13, there exists an urgent need for a standardized protocol for the isolation and culture of LLECs. Consequently, this study has successfully harvested and cultured LLECs by meticulously dissociating leptomeninges under the guidance of a microscope and promoting LLECs expansion through the use of vascular endothelial growth factor-C (VEGF-C). The distinctive marker for lymphatic endothelial cells is lymphatic vessel hyaluronic receptor-1 (LYVE-1)14. This multi-step protocol selectively isolates LYVE-1-positive LLECs using MACS and subsequently verifies their purity through flow cytometric analysis and immunofluorescent staining.
The primary steps of this multi-step protocol can be summarized as follows: flask coating, dissociation of leptomeninges, enzymatic digestion of leptomeninges, cell expansion, magnetic cell selection, and subsequent culture of LLECs. Finally, the purity of the isolated LLECs is confirmed through flow cytometric analysis and immunofluorescent staining. The overarching aim of this study is to present a reproducible, multi-step protocol for the isolation of LLECs from mouse leptomeninges and their subsequent in vitro culture. This protocol is poised to greatly facilitate investigations into the cellular functions and clinical implications of LLECs.

<table><tbody><tr><td>Block buffer</td><td>Beyotime</td><td>P0102</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>Collagenase I</td><td>Solarbio</td><td>C8140</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>DAPI</td><td>Beyotime</td><td>P0131</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>DMEM</td><td>Solarbio</td><td>11995</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>D-PBS</td><td>Solarbio</td><td>D1041</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>EGM-2 MV Bullet Kit</td><td>Lonza</td><td>C-3202</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>FBS</td><td>Solarbio</td><td>S9010</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>Fibronectin</td><td>Solarbio</td><td>F8180&amp;nbsp;</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>FlowJo Software</td><td>BD Biosciences</td><td>V10.8.1</td><td></td></tr><tr><td>LYVE-1 antibody</td><td>eBioscience</td><td>12-0443-82</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>Magnetic separator</td><td>Miltenyi</td><td>130-042-302</td><td>Sterile before use</td></tr><tr><td>Magnetic separator stand</td><td>Miltenyi</td><td>130-042-303</td><td>Sterile before use</td></tr><tr><td>Microbeads</td><td>Miltenyi</td><td>130-048-801</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>P/S</td><td>Solarbio</td><td>P1400</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>Papain</td><td>Solarbio</td><td>G8430-25g</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>PBS</td><td>Solarbio</td><td>D1040</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>PDPN antibody</td><td>Santa</td><td>sc-53533</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>PFA</td><td>Solarbio</td><td>P1110</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>PROX1 antibody</td><td>Santa</td><td>sc-81983</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr><tr><td>Selection column&amp;nbsp;</td><td>Miltenyi</td><td>130-042-401</td><td>Sterile before use</td></tr><tr><td>Trypsin</td><td>Gibco</td><td>25200072</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>VEGF-C&amp;nbsp;</td><td>Abcam</td><td>ab51947</td><td>Store aliquots at &amp;ndash;20 &amp;deg;C</td></tr><tr><td>VEGFR-3 antibody</td><td>Santa</td><td>sc-514825</td><td>Store aliquots at &amp;ndash;4 &amp;deg;C</td></tr></tbody></table>

harvest and primary culture of leptomeningeal lymphatic endothelial cells

Leptomeningeal lymphatic endothelial cells (LLECs) are a recently discovered intracranial cellular population with a unique distribution clearly distinct from peripheral lymphatic endothelial cells. Their cellular function and clinical implications remain largely unknown. Consequently, the availability of a supply of LLECs is essential for conducting functional research in vitro. However, there is currently no existing protocol for harvesting and culturing LLECs in vitro.
This study successfully harvested LLECs using a multi-step protocol, which included coating the flask with fibronectin, dissecting the leptomeninges with the assistance of a microscope, enzymatically digesting the leptomeninges to prepare a single-cell suspension, inducing the expansion of LLECs with vascular endothelial growth factor-C (VEGF-C), and selecting lymphatic vessel hyaluronic receptor-1 (LYVE-1) positive cells through magnetic-activated cell sorting (MACS). This process ultimately led to the establishment of a primary culture. The purity of the LLECs was confirmed through immunofluorescence staining and flow cytometric analysis, with a purity level exceeding 95%. This multi-step protocol has demonstrated reproducibility and feasibility, which will greatly facilitate the exploration of the cellular function and clinical implications of LLECs.

This study presents a reproducible, multi-step protocol for harvesting lymphatic endothelial cells (LLECs) from mice and subsequently establishing their primary culture in vitro. The key steps involve flask preparation and fibronectin coating, dissociation of leptomeninges, obtaining a single-cell suspension through enzymatic digestion, and inducing LLECs expansion with VEGF-C. LYVE-1-positive LLECs are then selectively isolated using magnetic-activated cell sorting (MACS). Finally, immunofluorescence staining a...

Watch this Scientific Journal Video about Unveiling Cellular Functions and Potential Clinical Implications of Leptomeningeal Lymphatic Endothelial Cells at JoVE.com

Author Spotlight: Unveiling Cellular Functions and Potential Clinical Implications of Leptomeningeal Lymphatic Endothelial Cells 

Leptomeningeal lymphatic endothelial cells (LLECs), a recently identified intracranial cell type, have poorly understood functions. This study presents a reproducible protocol for harvesting LLECs from mice and establishing in vitro primary cultures. This protocol is designed to enable researchers to delve into the cellular functions and potential clinical implications of LLECs.

Harvest and Primary Culture of Leptomeningeal Lymphatic Endothelial Cells

Leptomeningeal lymphatic endothelial cells (LLECs) are a recently discovered intracranial cellular population with a unique distribution clearly distinct ...

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Neuroscience

1.4K Views.  The First Affiliated Hospital of Kunming Medical University. Leptomeningeal lymphatic endothelial cells (LLECs), a recently identified intracranial cell type, have poorly understood functions. This study presents a reproducible protocol for harvesting LLECs from mice and establishing in vitro primary cultures. This protocol is designed to enable researchers to delve into the cellular functions and potential clinical implications of LLECs.

Video: Author Spotlight: Unveiling Cellular Functions and Potential Clinical Implications of Leptomeningeal Lymphatic Endothelial Cells 

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Murine Model for Studying Leptomeningeal Disease Using Circulating Tumor Cells

Ex Vivo Culture of Circulating Tumor Cells in the Cerebral Spinal Fluid from Melanoma Patients to Study Melanoma&#45;Associated Leptomeningeal Disease

Melanoma-associated leptomeningeal disease (M-LMD) occurs when circulating tumor cells (CTCs) enter into the cerebral spinal fluid (CSF) and colonize the meninges, the membrane layers that protect the brain and the spinal cord. Once established, the prognosis for M-LMD patients is dismal, with overall survival ranging from weeks to months. This is primarily due to a paucity in our understanding of the disease and, as a consequence, the availability of effective treatment options. Defining the underlying biology of M-LMD will significantly improve the ability to adapt available therapies for M-LMD treatment or design novel inhibitors for this universally fatal disease. A major barrier, however, lies in obtaining sufficient quantities of CTCs from the patient-derived CSF (CSF-CTCs) to conduct preclinical experiments, such as molecular characterization, functional analysis, and in vivo efficacy studies. Culturing CSF-CTCs ex vivo has also proven to be challenging. To address this, a novel protocol for the culture of patient-derived M-LMD CSF-CTCs ex vivo and in vivo is developed. The incorporation of conditioned media produced by human meningeal cells (HMCs) is found to be critical to the procedure. Cytokine array analysis reveals that factors produced by HMCs, such as insulin-like growth factor-binding proteins (IGFBPs) and vascular endothelial growth factor-A (VEGF-A), are important in supporting CSF-CTC survival ex vivo. Here, the usefulness of the isolated patient-derived CSF-CTC lines is demonstrated in determining the efficacy of inhibitors that target the insulin-like growth factor (IGF) and mitogen-activated protein kinase (MAPK) signaling pathways. In addition, the ability to intrathecally inoculate these cells in vivo to establish murine models of M-LMD that can be employed for preclinical testing of approved or novel therapies is shown. These tools can help unravel the underlying biology driving CSF-CTC establishment in the meninges and identify novel therapies to reduce the morbidity and mortality associated with M-LMD.

Author Spotlight: Assessing the Potential of Circulating Tumor Cells in Leptomeningeal Disease Research

This article describes a protocol for the propagation of cerebral spinal fluid-circulating tumor cells (CSF-CTCs) collected from patients with melanoma-associated leptomeningeal disease (M-LMD) to develop preclinical models to study M-LMD.

Melanoma-associated leptomeningeal disease (M-LMD) occurs when circulating tumor cells (CTCs) enter into the cerebral spinal fluid (CSF) and colonize the ...

Cancer Research

Isolation of Human Lymphatic Endothelial Cells by Multi-parameter Fluorescence-activated Cell Sorting

Lymphatic system disorders such as primary lymphedema, lymphatic malformations and lymphatic tumors are rare conditions that cause significant morbidity but little is known about their biology. Isolating highly pure human lymphatic endothelial cells (LECs) from diseased and healthy tissue would facilitate studies of the lymphatic endothelium at genetic, molecular and cellular levels. It is anticipated that these investigations may reveal targets for new therapies that may change the clinical management of these conditions. A protocol describing the isolation of human foreskin LECs and lymphatic malformation lymphatic endothelial cells (LM LECs) is presented. To obtain a single cell suspension tissue was minced and enzymatically treated using dispase II and collagenase II. The resulting single cell suspension was then labelled with antibodies to cluster of differentiation (CD) markers CD34, CD31, Vascular Endothelial Growth Factor-3 (VEGFR-3) and PODOPLANIN. Stained viable cells were sorted on a fluorescently activated cell sorter (FACS) to separate the CD34LowCD31PosVEGFR-3PosPODOPLANINPos LM LEC population from other endothelial and non-endothelial cells. The sorted LM LECs were cultured and expanded on fibronectin-coated flasks for further experimental use.

The goal of this protocol is to isolate lymphatic endothelial cells lining human lymphatic malformation cyst-like vessels and foreskins using fluorescence-activated cell sorting (FACS). Subsequent cell culturing and expansion of these cells permits a new level of experimental sophistication for genetic, proteomic, functional and cell differentiation studies.

Lymphatic system disorders such as primary lymphedema, lymphatic malformations and lymphatic tumors are rare conditions that cause significant morbidity ...

Medicine

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells (HUVECs) have shown to be convenient, easy to obtain and culture, and thus are the most widely studied endothelial cells. However, for research focused on processes like angiogenesis, permeability or many others, microvascular endothelial cells (ECs) are a much more physiologically relevant model to study 2. Furthermore, ECs isolated from knockout mice provide a useful tool for analysis of protein function ex vivo. Several approaches to isolate and culture microvascular ECs of different origin have been reported to date 3-7, but consistent isolation and culture of pure ECs is still a major technical problem in many laboratories. Here, we provide a step-by-step protocol on a reliable and relatively simple method of isolating and culturing mouse lung endothelial cells (MLECs). In this approach, lung tissue obtained from 6- to 8-day old pups is first cut into pieces, digested with collagenase/dispase (C/D) solution and dispersed mechanically into single-cell suspension. MLECS are purified from cell suspension using positive selection with anti-PECAM-1 antibody conjugated to Dynabeads using a Magnetic Particle Concentrator (MPC). Such purified cells are cultured on gelatin-coated tissue culture (TC) dishes until they become confluent. At that point, cells are further purified using Dynabeads coupled to anti-ICAM-2 antibody. MLECs obtained with this protocol exhibit a cobblestone phenotype, as visualized by phase-contrast light microscopy, and their endothelial phenotype has been confirmed using FACS analysis with anti-VE-cadherin 8 and anti-VEGFR2 9 antibodies and immunofluorescent staining of VE-cadherin. In our hands, this two-step isolation procedure consistently and reliably yields a pure population of MLECs, which can be further cultured. This method will enable researchers to take advantage of the growing number of knockout and transgenic mice to directly correlate in vivo studies with results of in vitro experiments performed on isolated MLECs and thus help to reveal molecular mechanisms of vascular phenotypes observed in vivo.

Here, we describe a protocol for isolation and culture of murine pulmonary endothelial cells. This method comprises mechanic and enzymatic lung tissue dissociation as well as a 2-step purification process using anti-PECAM-1 and anti-ICAM-2 antibodies conjugated to magnetic beads, which produces a pure endothelial cell population of mostly microvascular origin.

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells ...

Biology

Derivation of Leptomeninges Explant Cultures from Postmortem Human Brain Donors

Even though great progress has been made in the clinical characterization of Parkinson&#39;s disease, several studies report that the diagnosis of Parkinson&#39;s disease is not pathologically confirmed in up to 25% of clinically diagnosed Parkinson&#39;s disease. Therefore, tissue collected from clinically diagnosed patients with idiopathic Parkinson&#39;s disease can have a high rate of misdiagnosis; hence in vitro studies from such tissues to study Parkinson&#39;s disease as a preclinical model can become futile.
By collecting postmortem human leptomeninges with a confirmed neuropathological diagnosis of Parkinson&#39;s disease and characterized by nigrostriatal cell loss and intracellular protein inclusions called Lewy bodies, one can be certain that clinically observed parkinsonism is not caused by another underlying disease process (e.g. tumor, arteriosclerosis).
This protocol presents the dissection and preparation of postmortem human leptomeninges for derivation of a meningeal fibroblast culture. This procedure is robust and has a high success rate. The challenge of the culture is sterility as the brain procurement is generally not performed under sterile conditions. Therefore, it is important to supplement the culture media with a cocktail of penicillin, streptomycin, and amphotericin B.
The derivation of meningeal fibroblasts from autopsy-confirmed cases with Parkinson&#39;s disease provides the foundation for in vitro modeling of Parkinson&#39;s disease. Meningeal fibroblasts appear 3-9 days after sample preparation and about 20-30 million cells can be cryopreserved in 6-8 weeks. The meningeal fibroblast culture is homogenous and the cells express fibronectin, a commonly used marker to identify meninges.

The leptomeninges explant culture protocol from human postmortem brain is a technically robust and simple way to derive fibronectin-positive meningeal fibroblasts within 6-8 weeks and cryopreserve approximately 20-30 million cells.

Even though great progress has been made in the clinical characterization of Parkinson&#39;s disease, several studies report that the diagnosis of ...

Neisseria meningitidis Infection of Induced Pluripotent Stem-Cell Derived Brain Endothelial Cells

Meningococcal meningitis is a life-threatening infection that occurs when Neisseria meningitidis (meningococcus, Nm) can gain access to the central nervous system (CNS) by penetrating highly specialized brain endothelial cells (BECs). As Nm is a human-specific pathogen, the lack of robust in vivo model systems makes study of the host-pathogen interactions between Nm and BECs challenging and establishes a need for a human based model that mimics native BECs. BECs possess tighter barrier properties when compared to peripheral endothelial cells characterized by complex tight junctions and elevated trans-endothelial electrical resistance (TEER). However, many in vitro models, such as primary BECs and immortalized BECs, either lack or rapidly lose their barrier properties after removal from the native neural microenvironment. Recent advances in human stem-cell technologies have developed methods for deriving brain-like endothelial cells from induced pluripotent stem-cells (iPSCs) that better phenocopy BECs when compared to other in vitro human models. The use of iPSC-derived BECs (iPSC-BECs) to model Nm-BEC interaction has the benefit of using human cells that possess BEC barrier properties, and can be used to examine barrier destruction, innate immune activation, and bacterial interaction. Here we demonstrate how to derive iPSC-BECs from iPSCs in addition to bacterial preparation, infection, and sample collection for analysis.

Neisseria meningitidis Infection of Induced Pluripotent Stem-Cell Derived Brain Endothelial Cells

The protocol described here highlights the major steps in the differentiating induced pluripotent stem-cell derived brain-like endothelial cells, the preparation of Neisseria meningitidis for infection, and sample collection for other molecular analyses.

Meningococcal meningitis is a life-threatening infection that occurs when Neisseria meningitidis (meningococcus, Nm) can gain access to the central ...

Immunology and Infection

Induction of Leptomeningeal Cells Modification Via Intracisternal Injection

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the underlying parenchyma. Previously published protocols recommend using straight needles that should be lowered to a maximum of 1-2 mm from the dural surface. The sudden drop in resistance once the dural membrane has been punctured makes it difficult to maintain the needle in a steady position. Our method, instead, employs a needle bent at the tip that can be stabilized against the occipital bone of the skull, thus preventing the syringe from penetrating into the tissue after perforation of the dural membrane. The procedure is straightforward, reproducible, and does not cause long-lasting discomfort in the operated animals. We describe the intracisternal injection strategy in the context of genetic fate mapping of vascular leptomeningeal cells. The same technique can, furthermore, be utilized to address a wide range of research questions, such as probing the role of leptomeninges in neurodevelopment and the spreading of bacterial meningitis, through genetic ablation of genes putatively implicated in these phenomena. Additionally, the procedure can be combined with an automatized infusion system for a constant delivery and used for tracking cerebrospinal fluid movement via injection of fluorescently labelled molecules.

We describe an intracisternal injection that employs a needle bent at the tip that can be stabilized to the skull, thus eliminating the risk of damage to the underlying parenchyma. The approach can be used for genetic fate mapping and manipulations of leptomeningeal cells and for tracking cerebrospinal fluid movement.

The protocol outlined here describes how to safely and manually inject solutions through the cisterna magna while eliminating the risk of damage to the ...

Isolating Endothelial Cells from Murine Dermal Lymphatic Capillaries

Murine Dermal Lymphatic Endothelial Cell Isolation

The lymphatic system participates in the regulation of immune surveillance, lipid absorption, and tissue fluid balance. The isolation of murine lymphatic endothelial cells is an important process for lymphatic research, as it allows the performance of in vitro and biochemical experiments on the isolated cells. Moreover, the development of Cre-lox technology has enabled the tissue-specific deficiency of genes that cannot be globally targeted, leading to the precise determination of their role in the studied tissues. The dissection of the role of certain genes in lymphatic physiology and pathophysiology requires the use of lymphatic-specific promoters, and thus, the experimental verification of the expression levels of the targeted genes.
Methods for efficient isolation of lymphatic endothelial cells from wild-type or transgenic mice enable the use of ex vivo and in vitro assays to study the mechanisms regulating the lymphatic functions and the identification of the expression levels of the studied proteins. We have developed, standardized and present a protocol for the efficient isolation of murine dermal lymphatic endothelial cells (DLECs) via magnetic bead purification based on LYVE-1 expression. The protocol outlined aims to equip researchers with a tool to further understand and elucidate important players of lymphatic endothelial cell functions, especially in facilities where fluorescence-activated cell sorting equipment is not available.

Author Spotlight: Magnetic Bead-Based Isolation of Murine Dermal Lymphatic Endothelial Cells

This paper describes a method for magnetic bead-based isolation of murine endothelial cells from dermal lymphatic capillaries. The isolated lymphatic endothelial cells can be used for downstream in vitro experiments and protein expression analysis.

The lymphatic system participates in the regulation of immune surveillance, lipid absorption, and tissue fluid balance. The isolation of murine lymphatic ...

Studying Lymphocyte Transmigration Across a Microvascular Endothelial Cell Monolayer

Source: Schulte-Mecklenbeck, A., et al., <a href="https://app.jove.com/v/55390">Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier.</a> J. Vis. Exp. (2017)
This video demonstrates an experimental investigation of lymphocyte transmigration across a blood-brain barrier (BBB) endothelium model. A monolayer of human brain microvascular endothelial cells (HBMECs) is generated in vitro, and peripheral blood mononuclear cells (PBMCs) containing lymphocytes are added to the monolayer. The cells are then incubated to allow lymphocyte transmigration, and the migrated cells are quantified using flow cytometry.

Source: Schulte-Mecklenbeck, A., et al., Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier. J. Vis. Exp. ...

Harvest and Primary Culture of Leptomeningeal Lymphatic Endothelial Cells

Summary

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Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Ex Vivo Culture of Circulating Tumor Cells in the Cerebral Spinal Fluid from Melanoma Patients to Study Melanoma-Associated Leptomeningeal Disease

Isolation of Human Lymphatic Endothelial Cells by Multi-parameter Fluorescence-activated Cell Sorting

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Derivation of Leptomeninges Explant Cultures from Postmortem Human Brain Donors

Neisseria meningitidis Infection of Induced Pluripotent Stem-Cell Derived Brain Endothelial Cells

Induction of Leptomeningeal Cells Modification Via Intracisternal Injection

Murine Dermal Lymphatic Endothelial Cell Isolation

Studying Lymphocyte Transmigration Across a Microvascular Endothelial Cell Monolayer