We thank Steve McBryant and Jeff Hansen for help with ultracentrifugation and Patti Kiser for help with mouse handling. The National Institute of Neurological Diseases and Stroke at the National Institutes of Health, grant 5R01NS056379-02 funded this work.

1. Purifying and Labeling Prion Rods
This protocol is adapted from one previously published 18
<ol>
<li>Homogenize 100 grams of prion-infected brain tissue in 900 mL of ice cold homogenizing buffer (HB, 1X PBS containing 320 mM sucrose, 150 mM NaCl and 4mM EDTA) 1 min at maximum speed in a commercial blender, then 2 min on ice. Repeat three times.</li>
<li>Centrifuge homogenate for 10 min at 3000 x g and 4&deg;C. Remove and save supernatant on ice. Re-suspend pellet in 1L HB and repeat steps 1.1 and 1.2.</li>
<li>Pool supernatants and centrifuge for 60 min at 100,000 x g, 0&deg;C. Discard supernatant.</li>
<li>Re-suspend pellet in 100 mL of 1X Tris buffered saline (10 mM Tris-HCl, 0.1 mM NaCl and 1 mM EDTA) containing 2% Triton X-100 (TBST). Estimate protein concentration by the Bradford or similar assay and adjust to 5 mg/ mL in TBST. Chill on ice for 30 min.</li>
<li>Centrifuge for 30 min at 100,000 x g, 0&deg;C. Discard supernatant and wash pellet with 50 mL TBST then 100 mL TBS. Re-suspend pellet in 100 mL 1X PBS containing 1% sarcosyl and protease inhibitor cocktail and stir for 120 min at 37&deg;C.</li>
<li>Layer sample on 900 mL of 320 mM sucrose and centrifuge for 60 min at 100,000 x g, 10&deg;C. Discard supernatant and re-suspend pellet in 10 mL 2.3M NaCl, 5% sarcosyl.</li>
<li>Sonicate sample 5 x 40 seconds at 70% maximum power at 1 min intervals on ice.</li>
<li>Aliquot 1 mL of sample into eppendorf tubes and centrifuge for fifteen minutes at 13,000 x g, 4oC. Wash pellets twice with TBS and store @ -70&deg;C or conjugate to fluorochrome in step 1.9.</li>
<li>Conjugate 1 tube of purified prion rods using 1 vial of DyLight 649 fluorochrome according to manufacturer's protocol.</li>
<li>Exchange DyLight labeling buffer for PBS by dialysis or centrifugation through filter columns. Store in 20 &mu;L aliquots at -70&deg;C away from light until ready to use.</li>
</ol>
2. Prion Inoculation and Cell Recovery
This section encompasses intraperitoneal prion inoculation and recovery of cells from the peritoneum, mediastinal and mesenteric lymph nodes and spleen.
<ol>
<li>Dilute fluorescent prion rods 1:50 in PBS immediately prior to use. Scruff mouse by the dorsal neck fur with thumb and index finger, gently turn mouse to face you and restrain tail with pinky finger. Supinate mouse, elevating hind torso above head and holding him slightly upside down.</li>
<li>Using a 30G insulin syringe, inject 100 &mu;L of fluorescent prion rods 1 cm lateral of the midline and 1-2 cm anterior to the sacroiliac joint. Insert the needle 1 cm through the skin directed 45&deg; medially. Use 1:50 dilution of fluorescent beads or PBS alone as controls. Incubate mice for allotted time before analysis.</li>
<li>Euthanize mice according to approved animal care and use committee protocols. Inject 10 mL of PBS into the peritoneal cavity using a 12 mL syringe and 25G needle. Gently rock carcass longitudinally for one minute. Recover cell suspension from peritoneum using the same syringe and a 16G needle into a 15 mL conical tube and reserve on ice while carcass dissection continues.</li>
<li>Wet the ventral side of the carcass thoroughly with 70% ethanol. Lift the skin at the midline just below the ribcage with forceps and cut a small incision through the skin with scissors. Starting there, make a 3-5 cm long incision through the skin from in both directions to head and tail, then two 2-cm incisions from the midline laterally from each end. Peel back and pin the skin to reveal the peritoneal and thoracic cavities. Carefully cut through the thin translucent membranes surrounding the peritoneum and reflect the thorax to reveal the mediastinum.</li>
<li>Find and remove 2-4 mediastinal lymph nodes, slightly dorsal and laterally adjacent to the thymus, medially adjacent to the brachiocephalic artery, near the ventral side of the trachea. Find and remove up to six mesenteric lymph nodes, embedded in the soft white mesenteric tissue that anchors the intestines to the posterior abdominal wall. Find and remove the spleen, a long, dark red organ located just lateral of the midline and dorsal to the intestines on the left side of the abdomen. Reserve lymph nodes in 1 mL RPMI 1640 medium on ice until dissection is complete.</li>
<li>Macerate and press lymph nodes through a 40 &mu;m mesh strainer into 3 mL of RPMI in a plastic Petri dish using the plunger of a 1 mL syringe. Rinse strainer with an additional 2 mL of RPMI and transfer the 5 mL cell suspension to a 15 mL conical tube. Rinse the Petri dish with an additional 5 mL of RPMI and transfer to the same tube. Centrifuge 5 min at 250 x g, discard supernatant, re-suspend the cell pellet in 1 mL FACS buffer (1X PBS, 1% bovine serum albumin and 10 mM EDTA) and transfer to a 1.5 mL Eppendorf tube.</li>
</ol>
3. Cell Staining and Flow Cytometry
The following is a typical staining protocol for identifying immune cell subsets using fluorescent antibodies against well-characterized immune cell surface markers. All antibodies were diluted into FACS buffer immediately before use.
<ol>
<li>Count cells and aliquot 106 cells into Eppendorf tubes. Centrifuge cells (see 2.7) and wash pellet 2X with 1 mL FACS buffer.</li>
<li>Incubate cells 20 min on ice in 100 &mu;L of 2 ng/ mL rat anti-mouse CD16/32 to block endogenous Fc receptors. Pellet and wash cells with FACS buffer.</li>
<li>Add 100 &mu;L of 1ng/ mL appropriate fluorescent antibod(y)ies to each sample and incubate on ice for one hour. Add 100 &mu;L of FACS buffer alone to control cells. Pellet and wash cells twice with FACS buffer.</li>
<li>Re-suspend cells in 1 mL ACK buffer (150 mM NH4Cl, 1 mM KHCO3and 0.1 mM EDTA) Incubate on ice for 1 min to lyse red blood cells. Pellet and wash cells with FACS buffer. Re-suspend cells in 1 mL FACS buffer.</li>
<li>Collect data from cells using a flow cytometer capable of multicolor detection and analysis, typically with 405, 488 and 633 nm excitation lasers and five to ten fluorescence emission detectors.</li>
</ol>
4. Representative Results:
We purified prion rods from crude, prion-infected brain homogenate (Figure 1) Detergent solubilization and ultracentrifugation of the E2 elk brain homogenate (lanes 1 and 2) greatly enriched for prion rods (compare lanes 1 and 3), which was also protease K resistant (lane 4). Interestingly, undigested purified prion rods displayed a glycoform profile strikingly similar to digested E2 brain homogenate, indicating dramatic enrichment for prions. Identically-treated normal brain homogenate produced no PK-resistant PrPC (lanes 5 and 6).
Flow cytometric analysis of specific cell populations demonstrate that antigen presenting cells home to the peritoneum by two hours and retain fluorescent beads and prion rods, as evidenced by dramatically increased fluorescence over PBS injected controls (Figure 2). No cells harvested from the peritoneum of PBS-treated controls displayed fluorescence above background (Figs. 2A-G), while cells from bead (Figs. 2H-N) and prion (Figs. 2O-U)-inoculated mice displayed significant fluorescence, especially on monocytes (Figs. 2I and P), dendritic cells (Figs. 2K and R) and macrophages (Figs. 2L and S), and some Neutrophils (Figure 2J and Q) and fewer B cells (Figure 2T). Bead and Prion-bearing monocytes, dendritic cells and macrophages were also found in the mediastinal lymph nodes two hours after inoculation (Figs. 2AD and AK, AF and AM and AG and AN, respectively), while few or no PrPCWD-bearing Neutrophils, B or T cells were found there (Figs. 2AE and AL, AH and AG and AI and AP, respectively). We detected no beads or prion rods in the spleen or mesenteric lymph nodes (data not shown). In toto, these data demonstrate that immune cells traffic prions very similarly to other particulate antigens.
<table border="1">
<tbody>
 <tr>
 <td>REAGENT</td>
 <td>FLUOROCHROME</td>
 <td>EXCITATION LASER (nm)</td>
 <td>PEAK EMISSION (nm)</td>
 </tr>
 <tr>
 <td>Prion rods</td>
 <td>DyLight 649</td>
 <td>633</td>
 <td>674</td>
 </tr>
 <tr>
 <td>1 &mu;m beads</td>
 <td>AlexaFluor 660</td>
 <td>633</td>
 <td>685</td>
 </tr>
 <tr>
 <td>Antibodies</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CD11b</td>
 <td>eFluor 450</td>
 <td>405</td>
 <td>450</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>Phycoerythrin-Cy7</td>
 <td>488</td>
 <td>760</td>
 </tr>
 <tr>
 <td>CD11c</td>
 <td>R-Phycoerythrin</td>
 <td>488</td>
 <td>575</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>Phycoerythrin-Cy7</td>
 <td>488</td>
 <td>760</td>
 </tr>
 <tr>
 <td>Ly6C</td>
 <td>Fluoroisothiocyante</td>
 <td>488</td>
 <td>518</td>
 </tr>
 <tr>
 <td>Ly6G</td>
 <td>R-Phycoerythrin</td>
 <td>488</td>
 <td>575</td>
 </tr>
 <tr>
 <td>CD21</td>
 <td>DyLight 488</td>
 <td>488</td>
 <td>518</td>
 </tr>
 <tr>
 <td>B220</td>
 <td>Allophycocyanin-Cy7</td>
 <td>633</td>
 <td>785</td>
 </tr>
 <tr>
 <td>CD3</td>
 <td>R-Phycoerythrin</td>
 <td>488</td>
 <td>575</td>
 </tr>
</tbody>
</table>
Table 1. Spectral properties of fluorescent reagents 
 <img src="/files/ftp_upload/2349/2349fig1.jpg" alt="Figure 1" /> 
Figure 1. Purification of prion rods from infected brain homogenate. We used a 10% crude brain homogenate from a CWD-infected deer (E2, lanes 1and 2) as starting material from which we purified aggregated prion rods (lanes 3 and 4). Both crude and purified materials showed characteristic resistance to protease K treatment (lanes 1 - 4), whereas normal brain homogenate does not (lanes 5 and 6). Molecular weight markers are shown in Kd to the left of the blot. BH, brain homogenate.
 <img src="/files/ftp_upload/2349/2349fig2.jpg" alt="Figure 2" /> 
Figure 2. Flow cytometric analysis of immune cells trafficking prions from the peritoneum to mediastinal lymph nodes two hours after inoculation. PBS (panels A-G and V-AB), fluorescent beads (H-N and AC-AI) or Prion rods (O-U and AJ-AP) were injected into the peritoneum of mice and cells harvested from peritoneal lavage fluid (A-U) or mediastinal lymph nodes (V-AP) two hours later. Graphs in the first column show cells from mice treated with PBS (panels A and V), fluorescent beads (panel H and AC) and prion rods (panels O and AJ). Fluorescent cells (red or green dots) and total cells (grey dots) are plotted to show relative size (forward scatter), granularity (side scatter) and proportion of total live cells that fluoresce. Significant numbers of cells characteristic of granulocytes retained fluorescent beads and rods. Cells were also stained with antibodies against immune cell surface markers and gated for LyG- Ly6C+ monocytes (panels B, I, P, W, AD and AK), Ly6c- CD11c- CD11b+ Ly6G+ neutrophils (C, J, Q, X, AE and AL), Ly6G- Ly6C- CD11b+ CD11c+ dendritic cells (D, K, R, Y, AF and AM), Ly6G- Ly6C- CD11c- CD11b+ macrophages, (E, L, S, Z, AG and AN), CD3-B220+CD21+ B-cells (F, M, T, AA, AH and AO) and CD21-B220- CD3+ T cells (G, N, U, AB, AI and AP). Monocytes, dendritic cells and macrophages retained fluorescent beads and prion rods in the peritoneal cavity (panels I and P, K and R and L and S, respectively) and transported them to mediastinal lymph nodes (panels AD and AK, AF and AM and AG and AN, respectively), demonstrating similar uptake and trafficking of these two particulates. Neutrophils retained a significant amount of beads (J) and fewer rods (Q), but failed to deliver them to lymph nodes (AE and AL). B cells retained and transported some rods (T and AO) but virtually no beads (M and AH). T cells trafficked neither beads nor rods (N, U, AI and AP).

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporary+depletion+of+complement+component+C3+or+genetic+deficiency+of+C1q+significantly+delays+onset+of+scrapie.">Temporary depletion of complement component C3 or genetic deficiency of C1q significantly delays onset of scrapie.</a> Nat Med. 7, 485-485 (2001).</li><li>Zabel, M. 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No conflicts of interest declared.

Here we demonstrate a protocol for tagging and tracking prions in vivo that greatly facilitates monitoring early events in peripheral prion infection. This protocol greatly improves on past attempts at monitoring prion uptake in vitro 9 and in vivo 3,12 by pre-labeling highly enriched prion inoculum to unambiguously differentiate it from endogenous PrPC. Because infectious prions and PrPC share the same primary amino acid sequence, generating prion-...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>CWD-infected elk brain</td>
<td>&nbsp;</td>
<td>Private elk farm in Colorado</td>
<td>&nbsp;</td>
<td>Use any non-human prion-infected brain</td>
</tr>
<tr>
<td>Blender</td>
<td>&nbsp;</td>
<td>Oster</td>
<td>6694-015</td>
<td>Use any commercial blender</td>
</tr>
<tr>
<td>Centrifuge</td>
<td>&nbsp;</td>
<td>Sorvall</td>
<td>SS34 rotor</td>
<td>Use any centrifuge /rotor that can reach 3000 x g and hold &ge; 500 ml volumes</td>
</tr>
<tr>
<td>Ultracentrifuge</td>
<td>&nbsp;</td>
<td>Beckman</td>
<td>50.2 Ti rotor</td>
<td>Use any ultracentrifuge /rotor that can reach 100,000 x g and hold &ge; 500 ml volumes</td>
</tr>
<tr>
<td>Bradford Reagent</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>B6916</td>
<td>&nbsp;</td>
<tr>
<td>Complete mini protease inhibitor cocktail</td>
<td>&nbsp;</td>
<td>Roche</td>
<td>11 836 170 001</td>
<td>&nbsp;</td>
<tr>
<td>Sonicator</td>
<td>&nbsp;</td>
<td>Misonix</td>
<td>MP4000X</td>
<td>Use any horn or probe sonicator set to ~70% max power</td>
</tr>
<tr>
<td>DyLight antibody Labeling kit</td>
<td>&nbsp;</td>
<td>Thermo Scientific</td>
<td>53050</td>
<td>&nbsp;</td>
<tr>
<td>microcentrifuge</td>
<td>&nbsp;</td>
<td>Eppendorf</td>
<td>55430R</td>
<td>Use any refrigerated microcentrifuge that can achieve 13,000x g</td>
</tr>
<tr>
<td>centrifugal filter columns</td>
<td>&nbsp;</td>
<td>Millipore</td>
<td>Microcon YM-100</td>
<td>Use any filter or dialysis membrane with 100 Kd molecular weight cutoff</td>
</tr>
<tr>
<td>8-40 week-old FVB mice</td>
<td>&nbsp;</td>
<td>Charles River</td>
<td>207</td>
<td>Use any inbred mouse strain</td>
</tr>
<tr>
<td>1 &mu;m red fluorescent beads</td>
<td>&nbsp;</td>
<td>Phosphorex</td>
<td>2307</td>
<td>Use any fluorescent bead &le; 10 &mu;m</td>
</tr>
<tr>
<td>RPMI 1640 medium </td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>11875-093</td>
<td>&nbsp;</td>
<tr>
<td>40 &mu;m cell strainer</td>
<td>&nbsp;</td>
<td>Falcon</td>
<td>352340</td>
<td>&nbsp;</td>
<tr>
<td>fluorescent antibodies</td>
<td>&nbsp;</td>
<td>BD pharmingen</td>
<td>Various</td>
<td>Use any fluorescent antibody appropriate for your application.</td>
</tr>
<tr>
<td>flow cytometer</td>
<td>&nbsp;</td>
<td>Dakocytomation</td>
<td>CyanADP</td>
<td>Use any flow cytometer capable of multicolor fluorescence detection</td>
</tr>		</tbody>
		</table>
		</div>

monitoring immune cells trafficking fluorescent prion rods hours after intraperitoneal infection

Presence of an abnormal form a host-encoded prion protein (PrPC) that is protease resistant, pathologic and infectious characterizes prion diseases such as Chronic Wasting Disease (CWD) of cervids and scrapie in sheep. The Prion hypothesis asserts that this abnormal conformer constitutes most or all of the infectious prion. The role of the immune system in early events in peripheral prion pathogenesis has been convincingly demonstrated for CWD and scrapie 1-3. Transgenic and pharmacologic studies in mice revealed an important role of the Complement system in retaining and replicating prions early after infection 4-6. In vitro and in vivo studies have also observed prion retention by dendritic cells 7-10, although their role in trafficking remains unclear 11-16. Macrophages have similarly been implicated in early prion pathogenesis, but these studies have focused on events occurring weeks after infection 3,11,17. These prior studies also suffer from the problem of differentiating between endogenous PrPC and infectious prions. Here we describe a semiquantitative, unbiased approach for assessing prion uptake and trafficking from the inoculation site by immune cells recruited there. Aggregated prion rods were purified from infected brain homogenate by detergent solubilization of non-aggregated proteins and ultracentrifugation through a sucrose cushion. Polyacrylamide gel electrophoresis, coomassie blue staining and western blotting confirmed recovery of highly enriched prion rods in the pelleted fraction. Prion rods were fluorochrome-labeled then injected intraperitoneally into mice. Two hours later immune cells from peritoneal lavage fluid, spleen and mediastinal and mesenteric lymph nodes were assayed for prion rod retention and cell subsets identified by multicolor flow cytometry using markers for monocytes, neutrophils, dendritic cells, macrophages and B and T cells. This assay allows for the first time direct monitoring of immune cells acquiring and trafficking prions in vivo within hours after infection. This assay also clearly differentiates infectious, aggregated prions from PrPC normally expressed on host cells, which can be difficult and lead to data interpretation problems in other assay systems. This protocol can be adapted to other inoculation routes (oral, intravenous, intranervous and subcutaneous, e.g.) and antigens (conjugated beads, bacterial, viral and parasitic pathogens and proteins, egg) as well.

Watch this Scientific Journal Video about Monitoring Immune Cells Trafficking Fluorescent Prion Rods Hours after Intraperitoneal Infection at JoVE.com

Monitoring Immune Cells Trafficking Fluorescent Prion Rods Hours after Intraperitoneal Infection

Here we describe a novel assay for monitoring prion uptake and trafficking by immune cells immediately following intraperitoneal inoculation by purifying and fluorescently labeling aggregated prion rods from infected brain material then monitoring their uptake and movement from the injection site and characterizing the cells mediating these events.

Presence of an abnormal form a host-encoded prion protein (PrPC) that is protease resistant, pathologic and infectious characterizes prion diseases such ...

monitoring-immune-cells-trafficking-fluorescent-prion-rods-hours

Research

JoVE Journal

Immunology and Infection

11.4K visualizações.  Colorado State University. Aqui nós descrevemos um ensaio de romance para o monitoramento e captação de prion tráfico por células do sistema imunológico imediatamente após a inoculação intraperitoneal de purificação e fluorescente rotulagem varas prion agregados de material cerebral infectado acompanhamento da sua absorção e movimento a partir do local da injeção e caracterizar as células mediadoras desses eventos.

Vídeo: Monitoring Immune Cells Trafficking Fluorescent Prion Rods Hours after Intraperitoneal Infection

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

Seven Steps to Stellate Cells

Hepatic stellate cells are liver-resident cells of star-like morphology and are located in the space of Disse between liver sinusoidal endothelial cells and hepatocytes1,2. Stellate cells are derived from bone marrow precursors and store up to 80% of the total body vitamin A1, 2. Upon activation, stellate cells differentiate into myofibroblasts to produce extracellular matrix, thus contributing to liver fibrosis3. Based on their ability to contract, myofibroblastic stellate cells can regulate the vascular tone associated with portal hypertension4. Recently, we demonstrated that hepatic stellate cells are potent antigen presenting cells and can activate NKT cells as well as conventional T lymphocytes5. 

 Here we present a method for the efficient preparation of hepatic stellate cells from mouse liver. Due to their perisinusoidal localization, the isolation of hepatic stellate cells is a multi-step process. In order to render stellate cells accessible to isolation from the space of Disse, mouse livers are perfused in situ with the digestive enzymes Pronase E and Collagenase P. Following perfusion, the liver tissue is subjected to additional enzymatic treatment with Pronase E and Collagenase P in vitro. Subsequently, the method takes advantage of the massive amount of vitamin A-storing lipid droplets in hepatic stellate cells. This feature allows the separation of stellate cells from other hepatic cell types by centrifugation on an 8% Nycodenz gradient. The protocol described here yields a highly pure and homogenous population of stellate cells. Purity of preparations can be assessed by staining for the marker molecule glial fibrillary acidic protein (GFAP), prior to analysis by fluorescence microscopy or flow cytometry. Further, light microscopy reveals the unique appearance of star-shaped hepatic stellate cells that harbor high amounts of lipid droplets. 


 Taken together, we present a detailed protocol for the efficient isolation of hepatic stellate cells, including representative images of their morphological appearance and GFAP expression that help to define the stellate cell entity.

Here we describe a method for the isolation of hepatic stellate cells from mouse liver. For stellate cell purification, mouse livers are digested in situ and in vitro by pronase-collagenase treatment prior to density gradient centrifugation. This technique yields highly pure hepatic stellate cells.

Hepatic stellate cells are liver-resident  cells of star-like morphology and are located in the space of Disse between  liver sinusoidal endothelial cells ...

Isolation of Functional Cardiac Immune Cells

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5 These immune cells, which include mast cells, T-cells and macrophages; store and release a variety of biologically active mediators including cytokines and proteases such as tryptase.6-8 These mediators have been shown to be key players in extracellular matrix metabolism by activating matrix metalloproteinases or causing collagen accumulation by modulating the cardiac fibroblasts' function.9-11 However, available techniques for isolating cardiac immune cells have been problematic because they use bacterial collagenase to digest the myocardial tissue. This technique causes activation of the immune cells and thus a loss of function. For example, cardiac mast cells become significantly less responsive to compounds that cause degranulation.12 Therefore, we developed a technique that allows for the isolation of functional cardiac immune cells which would lead to a better understanding of the role of these cells in cardiac disease.13, 14
This method requires a familiarity with the anatomical location of the rat's xiphoid process, axilla and falciform ligament, and pericardium of the heart. These landmarks are important to increase success of the procedure and to ensure a higher yield of cardiac immune cells. These isolated cardiac immune cells can then be used for characterization of functionality, phenotype, maturity, and co-culture experiments with other cardiac cells to gain a better understanding of their interactions.

This method for isolating functional immune cells from the heart provides an alternative to the conventional methods of collagenase digestion, which causes unwanted immune cell activation, resulting in a decreased responsiveness of these cells. Our method of isolation yields functional cardiac immune cells by avoiding problems associated with enzymatic digestion.

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5  These immune cells, which ...

Following Cell-fate in E. coli After Infection by Phage Lambda

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the simultaneous infection of the cell by a number of phages, one of two pathways is chosen: lytic (virulent) or lysogenic (dormant)3,4. We recently developed a method for fluorescently labeling individual phages, and were able to examine the post-infection decision in real-time under the microscope, at the level of individual phages and cells5. Here, we describe the full procedure for performing the infection experiments described in our earlier work5. This includes the creation of fluorescent phages, infection of the cells, imaging under the microscope and data analysis. The fluorescent phage is a "hybrid", co-expressing wild- type and YFP-fusion versions of the capsid gpD protein. A crude phage lysate is first obtained by inducing a lysogen of the gpD-EYFP (Enhanced Yellow Fluorescent Protein) phage, harboring a plasmid expressing wild type gpD. A series of purification steps are then performed, followed by DAPI-labeling and imaging under the microscope. This is done in order to verify the uniformity, DNA packaging efficiency, fluorescence signal and structural stability of the phage stock. The initial adsorption of phages to bacteria is performed on ice, then followed by a short incubation at 35&deg;C to trigger viral DNA injection6. The phage/bacteria mixture is then moved to the surface of a thin nutrient agar slab, covered with a coverslip and imaged under an epifluorescence microscope. The post-infection process is followed for 4 hr, at 10 min interval. Multiple stage positions are tracked such that ~100 cell infections can be traced in a single experiment. At each position and time point, images are acquired in the phase-contrast and red and green fluorescent channels. The phase-contrast image is used later for automated cell recognition while the fluorescent channels are used to characterize the infection outcome: production of new fluorescent phages (green) followed by cell lysis, or expression of lysogeny factors (red) followed by resumed cell growth and division. The acquired time-lapse movies are processed using a combination of manual and automated methods. Data analysis results in the identification of infection parameters for each infection event (e.g. number and positions of infecting phages) as well as infection outcome (lysis/lysogeny). Additional parameters can be extracted if desired.

Following Cell-fate in E. coli After Infection by Phage Lambda

This article describes the procedure for preparing a fluorescently-labeled version of bacteriophage lambda, infection of E. coli bacteria, following the infection outcome under the microscope, and analysis of infection results.

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the ...

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Throughout the last years, the contribution of alveolar type II epithelial cells (AECII) to various aspects of immune regulation in the lung has been increasingly recognized. AECII have been shown to participate in cytokine production in inflamed airways and to even act as antigen-presenting cells in both infection and T-cell mediated autoimmunity 1-8. Therefore, they are especially interesting also in clinical contexts such as airway hyper-reactivity to foreign and self-antigens as well as infections that directly or indirectly target AECII. However, our understanding of the detailed immunologic functions served by alveolar type II epithelial cells in the healthy lung as well as in inflammation remains fragmentary. Many studies regarding AECII function are performed using mouse or human alveolar epithelial cell lines 9-12. Working with cell lines certainly offers a range of benefits, such as the availability of large numbers of cells for extensive analyses. However, we believe the use of primary murine AECII allows a better understanding of the role of this cell type in complex processes like infection or autoimmune inflammation. Primary murine AECII can be isolated directly from animals suffering from such respiratory conditions, meaning they have been subject to all additional extrinsic factors playing a role in the analyzed setting. As an example, viable AECII can be isolated from mice intranasally infected with influenza A virus, which primarily targets these cells for replication 13. Importantly, through ex vivo infection of AECII isolated from healthy mice, studies of the cellular responses mounted upon infection can be further extended.
Our protocol for the isolation of primary murine AECII is based on enzymatic digestion of the mouse lung followed by labeling of the resulting cell suspension with antibodies specific for CD11c, CD11b, F4/80, CD19, CD45 and CD16/CD32. Granular AECII are then identified as the unlabeled and sideward scatter high (SSChigh) cell population and are separated by fluorescence activated cell sorting 3.
In comparison to alternative methods of isolating primary epithelial cells from mouse lungs, our protocol for flow cytometric isolation of AECII by negative selection yields untouched, highly viable and pure AECII in relatively short time. Additionally, and in contrast to conventional methods of isolation by panning and depletion of lymphocytes via binding of antibody-coupled magnetic beads 14, 15, flow cytometric cell-sorting allows discrimination by means of cell size and granularity. Given that instrumentation for flow cytometric cell sorting is available, the described procedure can be applied at relatively low costs. Next to standard antibodies and enzymes for lung disintegration, no additional reagents such as magnetic beads are required. The isolated cells are suitable for a wide range of functional and molecular studies, which include in vitro culture and T-cell stimulation assays as well as transcriptome, proteome or secretome analyses 3, 4.

We describe the rapid isolation of primary murine type II alveolar epithelial cells (AECII) by flow cytometric negative selection. These AECII show high viability and purity and are suitable for a wide range of functional and molecular studies regarding their role in respiratory conditions such as autoimmune or infectious diseases.

Throughout  the last years, the contribution of alveolar type II epithelial cells (AECII)  to various aspects of immune regulation in the lung has been ...

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 ...

Generation of Lymph Node-fat Pad Chimeras for the Study of Lymph Node Stromal Cell Origin

The stroma is a key component of the lymph node structure and function. However, little is known about its origin, exact cellular composition and the mechanisms governing its formation. Lymph nodes are always encapsulated in adipose tissue and we recently demonstrated the importance of this relation for the formation of lymph node stroma. Adipocyte precursor cells migrate into the lymph node during its development and upon engagement of the Lymphotoxin-b receptor switch off adipogenesis and differentiate into lymphoid stromal cells (B&#233;n&#233;zech&#160;et al.14). Based on the lymphoid stroma potential of adipose tissue, we present a method using a lymph node/fat pad chimera that allows the lineage tracing of lymph node stromal cell precursors. We show how to isolate newborn lymph nodes and EYFP+ embryonic adipose tissue and make a LN/ EYFP+ fat pad chimera. After transfer under the kidney capsule of a host mouse, the lymph node incorporates local adipose tissue precursor cells and finishes its formation. Progeny analysis of EYFP+ fat pad cells in the resulting lymph nodes can be performed by flow-cytometric analysis of enzymatically digested lymph nodes or by immunofluorescence analysis of lymph nodes cryosections. By using fat pads from different knockout mouse models, this method will provide an efficient way of analyzing the origin of the different lymph node stromal cell populations.

Generation of lymph node/fat pad chimeras for the study of lymph node stromal cell origin is described. The method involves the isolation of lymph nodes from newborn mice and embryonic fat pads, the generation of chimeric lymph node-fat pads, and their transfer under the kidney capsule of a host mouse.

The stroma is a key component of the lymph node structure and function. However, little is known about its origin, exact cellular composition and the ...

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact cellular functions, including energetics and signal transductions. While both fluorescent and electrophysiological methods, including electrode usage and patch-clamping, have been well developed for measuring these events in eukaryotic cells, methodology for measuring similar events in microorganisms have proven more challenging to develop given their small size in combination with the more complex outer surface of bacteria shielding the membrane. During our studies of death-initiation in Streptococcus pneumoniae (pneumococcus), we wanted to elucidate the role of membrane events, including changes in polarity, integrity, and intracellular ion concentrations. Searching the literature, we found that very few studies exist. Other investigators had monitored radioisotope uptake or equilibrium to measure ion fluxes and membrane potential and a limited number of studies, mostly in Gram-negative organisms, had seen some success using carbocyanine or oxonol fluorescent dyes to measure membrane potential, or loading bacteria with cell-permeant acetoxymethyl (AM) ester versions of ion-sensitive fluorescent indicator dyes. We therefore established and optimized protocols for measuring membrane potential, rupture, and ion-transport in the Gram-positive organism S. pneumoniae. We developed protocols using the bis-oxonol dye DiBAC4(3) and the cell-impermeant dye propidium iodide to measure membrane depolarization and rupture, respectively, as well as methods to optimally load the pneumococci with the AM esters of the ratiometric dyes Fura-2, PBFI, and BCECF to detect changes in intracellular concentrations of Ca2+, K+, and H+, respectively, using a fluorescence-detection plate reader. These protocols are the first of their kind for the pneumococcus and the majority of these dyes have not been used in any other bacterial species. Though our protocols have been optimized for S. pneumoniae, we believe these approaches should form an excellent starting-point for similar studies in other bacterial species.

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Unlike that seen for eukaryotes, there is a paucity of studies that detail membrane depolarization and ion concentration changes in bacteria, primarily as their small size makes conventional methods of measurement difficult. Here, we detail protocols for monitoring such events in the significant Gram-positive pathogen Streptococcus pneumoniae utilizing fluorescence techniques.

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact ...

Detection of Fluorescent Nanoparticle Interactions with Primary Immune Cell Subpopulations by Flow Cytometry

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable method of analysis by flow cytometry to study nanoparticle interaction with immune cells. Primary immune cells can be easily purified from human or mouse tissues by antibody-mediated magnetic isolation. In the first instance, the different cell populations running in a flow cytometer can be distinguished by the forward-scattered light (FSC), which is proportional to cell size, and the side-scattered light (SSC), related to cell internal complexity. Furthermore, fluorescently labeled antibodies against specific cell surface receptors permit the identification of several subpopulations within the same sample. Often, all these features vary when cells are boosted by external stimuli that change their physiological and morphological state. Here, 50 nm FITC-SiO2 nanoparticles are used as a model to identify the internalization of nanostructured materials in human blood immune cells. The cell fluorescence and side-scattered light increase after incubation with nanoparticles allowed us to define time and concentration dependence of nanoparticle-cell interaction. Moreover, such protocol can be extended to investigate Rhodamine-SiO2 nanoparticle interaction with primary microglia, the central nervous system resident immune cells, isolated from mutant mice that specifically express the Green Fluorescent Protein (GFP) in the monocyte/macrophage lineage. Finally, flow cytometry data related to nanoparticle internalization into the cells have been confirmed by confocal microscopy.

Analysis of nanoparticle interaction with defined subpopulations of immune cells by flow cytometry.

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable ...

A Semi-automated Approach to Preparing Antibody Cocktails for Immunophenotypic Analysis of Human Peripheral Blood

Immunophenotyping of peripheral blood by flow cytometry determines changes in the frequency and activation status of peripheral leukocytes during disease and treatment. It has the potential to predict therapeutic efficacy and identify novel therapeutic targets. Whole blood staining utilizes unmanipulated blood, which minimizes artifacts that can occur during sample preparation. However, whole blood staining must also be done on freshly collected blood to ensure the integrity of the sample. Additionally, it is best to prepare antibody cocktails on the same day to avoid potential instability of tandem-dyes and prevent reagent interaction between brilliant violet dyes. Therefore, whole blood staining requires careful standardization to control for intra and inter-experimental variability.
Here, we report deployment of an automated liquid handler equipped with a two-dimensional (2D) barcode reader into a standard process of making antibody cocktails for flow cytometry. Antibodies were transferred into 2D barcoded tubes arranged in a 96 well format and their contents compiled in a database. The liquid handler could then locate the source antibody vials by referencing antibody names within the database. Our method eliminated tedious coordination for positioning of source antibody tubes. It provided versatility allowing the user to easily change any number of details in the antibody dispensing process such as specific antibody to use, volume, and destination by modifying the database without rewriting the scripting in the software method for each assay.
A proof of concept experiment achieved outstanding inter and intra- assay precision, demonstrated by replicate preparation of an 11-color, 17-antibody flow cytometry assay. These methodologies increased overall throughput for flow cytometry assays and facilitated daily preparation of the complex antibody cocktails required for the detailed phenotypic characterization of freshly collected anticoagulated peripheral blood.

Whole blood Immunophenotyping is indispensable for monitoring the human immune system. However, the number of reagents required and the instability of specific fluorochromes in premixed cocktails necessitates daily reagent preparation. Here we show that semi-automated preparation of staining antibody cocktail helps establish reliable immunophenotyping by minimizing variability in reagent dispensing.

Immunophenotyping of peripheral blood by flow cytometry determines changes in the frequency and activation status of peripheral leukocytes during disease ...