NGL was supported by the Wayne State University Perinatal Initiative in Maternal, Perinatal and Child Health. We gratefully acknowledge Maureen McGerty and Amy E. Furcron (Wayne State University) for their critical reading of the manuscript.

Before working with the samples mentioned in this protocol, animal ethical approval must be given by the Local Research Ethics Committee and the Institutional Review Boards. When working with animal blood, cells, or hazardous agents as mentioned in this protocol, the proper biosafety and laboratory safety actions must be followed.
1. Mouse Handling and Tissue Collection
<ol>
	<li>Prepare a sterile workstation and obtain sterile tools for tissue collection. These tools will include large and small surgical scissors, forceps, and fine-tip tweezers. Include small Petri dishes labeled with the appropriate tissue names, filled with 1x phosphate-buffered saline (PBS) solution (3 - 5 ml) equilibrated to RT (20 - 22 &#176;C). Also include a large Petri dish, unlabeled, filled with 1x PBS solution (10 - 15 ml).</li>
	<li>Euthanize a pregnant mouse (10.5 to 19.0 days post-coitum (dpc) or prior to delivery) using carbon dioxide (CO2). To ensure that the mouse is euthanized, use the cervical dislocation technique. 
	Note: Prior to 10.5 dpc, it is difficult to manually separate the decidual tissues from the uterine tissues. If the isolation of decidual leukocytes is not required, leukocytes from the uterine tissues of non-pregnant mice or those from mice at &#60;10.5 dpc can also be performed by following this protocol.</li>
	<li>Using a pair of sterile surgical scissors, completely remove the lower abdominal portion of the skin and muscular tissue. With forceps, move aside all other organs until the uterus and ovaries are visible. With the uterus still attached, use the forceps to move it outside of the peritoneal cavity (Figure 1A).</li>
	<li>Locate the ovaries distal to the oviduct and uterotubal junction of each uterine horn. Using the small surgical scissors, make an incision at the uterotubal junction and separate the uterine horns from the mesentery containing the uterine vessels; then, excise at the cervix to detach the uterine horns from the peritoneal cavity (Figure 1B).</li>
	<li>With small surgical scissors, make an incision between the cervix and the lower segment of the uterine horns. Leave attached a portion of the cervix during this process (Figure 1C) to avoid opening the uterine horn.</li>
	<li>Immerse the uterine horns (including the cervix) in a large Petri dish filled with 1x PBS solution (10 - 15 ml) to hydrate the implantation sites.</li>
	<li>While still immersed in 1x PBS solution, trim any fat from the uterine horns with the small surgical scissors. Keep the tissues immersed in 1x PBS solution throughout the dissection.</li>
	<li>Holding the uterine horns in place with the forceps, remove one of the implantation sites from the uterus with a transverse cut through the inter-implantation region, using the small surgical scissors (Figure 1D). The implantation site contains the fetus, surrounded by the chorioallantoic membrane, and the uterine, decidual, and placental tissues.</li>
	<li>Using the forceps, hold the site in place and make a small incision in the uterine wall adjacent to the placental and decidual tissues (Figure 1E). Trim around the perimeter of the placenta/decidua until these become separated from both the uterine wall and the chorioallantoic membrane that includes the fetus (Figure 1F).</li>
	<li>Remove the placenta and attached decidual tissues and place in 1x PBS solution (3 - 5&#160; ml; see step 1.12).</li>
	<li>Place the fetus surrounded by the chorioallantoic membrane attached to the uterine tissues in a Petri dish filled with 1x PBS solution (3 - 5&#160; ml). The hydration of these tissues will facilitate the separation of the chorioallantoic membrane from the uterine wall (see step 1.14).</li>
	<li>Using a pair of fine-tip tweezers, gently peel the decidual tissue (white-grey layer) from the placental surface. Perform this step carefully to maintain the integrity of the placental and decidual tissues (Figure 1G).</li>
	<li>Place the placenta and decidual tissues in two separately labeled Petri dishes filled with 1x PBS solution (3 - 5&#160; ml each).</li>
	<li>Gently remove the uterine wall from the chorioallantoic membrane with fine-tip tweezers. These are the uterine tissues.</li>
	<li>Place the uterine tissues in a Petri dish filled with 1x PBS solution (3 - 5&#160; ml).</li>
	<li>Repeat steps 1.8 through 1.15 until all implantation sites have been processed.</li>
	<li>Collect several uterine, decidual, and placental tissues to improve the yield of cells during enzymatic digestion. The amount of tissues depends on the litter size; however, leukocyte isolation requires &#62;2 pieces of tissue. 
	Note: For more detailed information regarding the dissection of the implantation sites on different gestational days, view the pictures found in Chapter Two of The Guide to Investigation of Mouse Pregnancy42.</li>
</ol>
2. Uterine, Decidual, and Placental Tissue Dissociation
<ol>
	<li>Label several sterile 2 ml conical tubes with the appropriate tissue name, and place a vial with 1,000 &#181;l&#160;of enzymatic solution on ice.</li>
	<li>From the Petri dish filled with 1x PBS solution, transfer two pieces (100 - 150 mg) of the uterine, decidual, or placental tissues to a labeled 2 ml conical tube.</li>
	<li>Add 500 &#181;l of cold enzymatic solution into each 2 ml conical tube.</li>
	<li>With fine sterile scissors, begin to dissociate the tissue immersed in the enzymatic solution until it is finely minced. Over time, the suspension will develop a milky appearance. This step should not exceed more than 2 min to prevent over-manipulation of the sample.</li>
	<li>Once the tissue has been dissociated, add an extra 500 &#181;l of cold enzymatic solution to the 2 ml conical tube.</li>
	<li>Place the 2 ml conical tube on ice.</li>
	<li>Repeat steps 2.2 through 2.6 with each individual tissue until all tissues have been processed.</li>
	<li>Transfer all 2 ml conical tubes with homogenized tissue (minced tissue + enzymatic solution) samples from the ice into an incubator at 37&#160;&#186;C and perform a gentle orbital agitation (80 rpm). Incubate for 30 - 35 min. Make sure the temperature of the incubator is stabilized before starting the incubation time.</li>
	<li>Following incubation, remove the 2 ml conical tubes with dissociated tissues from the incubator and place them on ice to prevent any further digestion.</li>
	<li>Label sterile 50 ml conical tubes according to tissue type. Remove the lids and replace them with a sterile cell strainer (100 &#181;m pore size).</li>
	<li>Pour the dissociated tissues from the 2 ml conical tubes through the cell strainer using ~20 ml of 1x PBS solution. The fragments of tissue will remain in the cell strainer and the cell suspension will pass through it.</li>
	<li>Wash the dissociated tissues in the cell strainer 2 - 3 times with 20 ml of 1x PBS solution using a transfer pipette.</li>
	<li>Rinse the empty 2 ml conical tubes with 1x PBS solution (1 ml) and pour the contents through the cell strainers.</li>
	<li>Remove the cell strainers from the 50 ml conical tubes and replace the lids. Centrifuge the tubes at 1,250 x g for 10 min at 4 &#186;C.</li>
	<li>Carefully, aspirate the supernatant without disturbing the cell pellet. The cell pellet contains the isolated leukocytes and stromal cells.</li>
	<li>Re-suspend the cell pellet in 1 ml of RPMI culture medium without fetal bovine serum (FBS). Mix gently. Do not use vortex.</li>
	<li>Add 500 &#181;l of neat FBS to a 5 ml polystyrene plastic tube and slowly overlay the cell suspension.</li>
	<li>Centrifuge for 10 min at 1,100 x g without the brake at RT to pellet viable cells while cellular debris is retained at the PBS/FBS interface.</li>
	<li>Aspirate the supernatant without touching the cell pellet. The cell pellet contains mostly viable cells.</li>
	<li>Optionally: To concentrate the mononuclear cells, use a 20% saturated polysaccharide solution, as described below.
	<ol>
		<li>Resuspend the cell pellet in 1,000 &#181;l of RPMI culture medium without FBS.</li>
		<li>Add 1 ml of 20% saturated polysaccharide solution to a 5 ml polystyrene plastic tube and slowly overlay the cell suspension on top of this solution, according to the manufacturer&#8217;s instructions. While layering the sample, avoid mixing the 20% saturated polysaccharide solution with the cell suspension.</li>
		<li>Centrifuge with a swing-out rotor at 500 x g for 30 min at RT without brake. It is very important to turn off the brake to avoid mixing the cells and losing the gradient. Mononuclear cells will be found in the interface between the 20% saturated polysaccharide solution and the 1x PBS solution.</li>
		<li>Aspirate and discard the supernatant. The cell pellet contains mostly mononuclear cells. 
		Note: To perform viability staining and intracellular staining, see step 3. To perform immunophenotyping, see step 4. To perform viability staining and extracellular staining only, see step 5. To perform a cell culture of isolated leukocytes, see step 6. To perform magnetic cell sorting, see step 7.</li>
	</ol>
	</li>
</ol>
3. Viability Staining for Fixable Cells
<ol>
	<li>Resuspend the cell pellet in 1,100 &#181;l of 1x PBS solution. Mix gently using a micro-pipette.</li>
	<li>Transfer 100 &#181;l of the cell suspension into a 5 ml polystyrene plastic tube. Add 400 &#181;l of 1x PBS solution to the 5 ml polystyrene tube. This tube is a control for tissue auto-fluorescence. Store at 4 &#186;C until step 3.5.</li>
	<li>Transfer the remaining 1,000 &#181;l of the cell suspension to a new 5 ml polystyrene plastic tube. Add 1 &#181;l of the viability dye and gently homogenize. Incubate for 30 min in darkness at 4 &#186;C.</li>
	<li>Following incubation, add 1,000 &#181;l of 1x PBS solution. Mix gently by hand.</li>
	<li>Centrifuge both 5 ml polystyrene plastic tubes (steps 3.2 and 3.3) at 1250 x g for 10 min at 4 &#186;C. Aspirate and discard the supernatant. 
	Note: The cell pellet (step 3.3) will be used in step 4. The cell pellet (step 3.2) will serve as a control for tissue auto-fluorescence.</li>
</ol>
4. Immunophenotyping
<ol>
	<li>Resuspend the cell pellet in 50 &#181;l of anti-mouse CD16/CD32 (diluted 1:100 in FACS buffer [0.1% BSA, 0.05% sodium azide, 1x PBS solution, ph 7.4]). Mix the cell suspension gently. Do not use the vortex.</li>
	<li>Incubate the cell suspension in the 5 ml polystyrene tube in darkness at 4 &#176;C for 10 min.</li>
	<li>Following incubation, add 50 &#181;l of each anti-leukocyte monoclonal antibody reacting with extracellular cell surface markers or isotype-matched control antibody (antibody dilutions must be validated). For example, to analyze leukocyte sub-populations, use anti-mouse antibodies reacting with the following leukocyte-cell surface markers: CD45, Ly6G, F4/80, CD11c, CD49b, B220, CD3, CD4, and CD8 (Table 1).</li>
	<li>Once antibodies against extracellular markers have been added to the 5 ml polystyrene plastic tube, mix gently. Incubate in darkness for 30 min at 4 &#176;C.</li>
	<li>During this incubation, prepare and pre-warm the fixation buffer solution (diluted with deionized water, 1:5) to 37 &#186;C, if required. Keep the buffer at 37 &#186;C in darkness until its use is necessary.</li>
	<li>After 30 min incubation, add 500 &#181;l of FACS buffer in the 5 ml polystyrene plastic tube and gently mix. This step is needed to wash the cells and remove any excess of unbound antibody.</li>
	<li>Centrifuge the samples at 1,250 x g at 4 &#176;C for 10 min. Aspirate and discard the supernatant. 
	Note: If intracellular staining is not required, see step 5. If intracellular staining is required, perform permeabilization and intracellular staining here, and then proceed with the fixation (step 4.8).</li>
	<li>Add 500 &#181;l of the pre-warmed fixation buffer solution to the cell pellet and incubate in darkness for 10 min at 37 &#186;C.</li>
	<li>Dispense 500 &#181;l of FACS buffer into the 5 ml polystyrene plastic tube. Mix gently.</li>
	<li>Centrifuge the samples at 1,250 x g at 4 &#176;C for 10 min. Aspirate and discard the supernatant.</li>
	<li>Add 500 &#181;l of FACS buffer to each sample. These samples can now be analyzed by flow cytometry. Acquire the samples immediately for best results.</li>
</ol>
5. Viability Staining for Unfixable Cells
<ol>
	<li>After extracellular immunophenotyping, resuspend the cell pellet in 500 &#181;l of FACS buffer.</li>
	<li>Add 1 &#181;l of 4&#8217;,6-diamidino-2-phenylindole dihydrochloride (DAPI, 200 &#181;g/ml) just before acquiring the sample.</li>
	<li>Visualize the samples within 1 - 2 min after adding DAPI using a flow cytometer.</li>
</ol>
6. Cell Culture
<ol>
	<li>Prepare a workstation under a sterile fume hood. Pre-warm the RPMI culture medium to 37 &#176;C using a water bath.</li>
	<li>Resuspend the cell pellet in 1000 &#181;l of FACS buffer for cell counting.</li>
	<li>Count the number of viable cells using an automatic cell counter or hemocytometer and 0.4% trypan blue solution, following the manufacturer&#39;s instructions. Record the total live cell count.</li>
	<li>Centrifuge the cells at 1,250 x g at 4 &#176;C for 10 min to remove the FACS buffer.</li>
	<li>Resuspend the cell pellet in 500 &#181;l of pre-warmed RPMI culture medium by gently pipetting 2 to 3 times.</li>
	<li>Plate the desired number of cells in a sterile 24-well plate and incubate at 37 &#176;C. The incubation time will be determined by the research question.</li>
</ol>
7. Magnetic Cell Sorting
<ol>
	<li>Set up a clean workstation with the following materials: MS columns, magnetic cell separator, 15 ml conical tubes, 30 &#181;m pre-separation filters, and a multi-stand.</li>
	<li>Resuspend the cell pellet in 500 &#181;l of MACS buffer (0.5% BSA, 2mM EDTA, 1x PBS solution, pH 7.2).</li>
	<li>Using an automatic cell counter or hemocytometer and 0.4% trypan blue solution, count and record the total live cell count.</li>
	<li>Centrifuge the cells at 1,250 x g at 4 &#176;C for 10 min. Aspirate and discard the supernatant. Add another 500 &#181;l of MACS buffer.</li>
	<li>Proceed to perform the magnetic separation following the manufacturer&#8217;s recommendations.</li>
	<li>Determine purity by flow cytometry or microscopy.</li>
</ol>

The authors disclose no conflicts of interest.

The collection of consistent data that records the abundance and phenotypic characteristics of infiltrating leukocytes at the maternal-fetal interface is essential to understanding the pathogenesis of pregnancy-related complications. Several techniques have been described that facilitate the isolation of infiltrating leukocytes from the murine tissues at the maternal-fetal interface throughout pregnancy31,38,39,43-46. However, each technique is different, uses different enzymes or enzyme combinations, requires...

Immune tolerance in pregnancy is a period when distinctive changes occur within the immune system of the mother. These changes allow the mother to tolerate the fetus, a semi-allogenic graft1. The fetus expresses paternal major histocompatibility complex (MHC) antigens2, and fetal cells have been found in the maternal circulation3; however, the fetus is not rejected4,5. This enigma is not fully understood.
The most recent hypothesis states that maternal-fetal tolerance is created during coitus and fecundation6,7 and maintained to sustain a full-term pregnancy8-10. A breakdown of this maternal-fetal tolerance is considered a mechanism of disease during early and late stages of pregnancy10-16. Maternal-fetal tolerance involves the participation of various leukocyte sub-populations, including T cells (regulatory T cells, Th1 cells, Th2 cells, and Th17 cells), macrophages, neutrophils, mast cells, NK cells, and NKT cells, dendritic cells, and B cells, that change in density and localization throughout pregnancy15,17-19. Maternal-fetal tolerance is enriched at the maternal-fetal interface20 - the anatomical site where the immune system of the mother interacts with the fetal antigens20,21.
The maternal-fetal interface is created during placentation when the fetal extravillous trophoblast cells invade the uterine mucosa22-24. On the fetal side of this interface, the membranes surrounding the fetus create a specialized epithelial surface within the placenta, and the syncytiotrophoblast cells control the nutrient exchange through their direct contact with maternal blood22. On the maternal side of the interface, the decidua recruits a heterogeneous pool of leukocytes that in mice account for 30% to 50% of all decidual cells. In addition to their participation in maternal immune tolerance, these cells are key contributors to different processes during pregnancy, e.g., the protection of the reproductive tract from infections, fecundation, embryo implantation7,25, decidual angiogenesis26, vascular remodeling24,27, trophoblast invasion28, placental development24,25, and, ultimately, labor and delivery15,17. Therefore, the study of the leukocytes involved in maternal-fetal tolerance is essential to elucidating the pathogenesis of pregnancy-related complications.
While the use of immunohistochemistry and immunofluorescence has generated data for the direct visualization and localization of uterine, decidual, or placental leukocytes29,30, flow cytometry analysis has further revealed specific subsets of leukocytes in each of these tissues31,32. Additionally, flow cytometry has been used to determine the density and proportion of maternal-fetal interface leukocytes33 and expression levels of extracellular and intracellular proteins8-10,34. Flow cytometric analysis of leukocytes at the maternal-fetal interface requires a single-cell suspension. In order to isolate infiltrating leukocytes from the decidual, uterine, and placental tissues, two methods of tissue dissociation have been used: mechanical and enzymatic. Both methods allow the separation of infiltrated leukocytes from the extracellular matrix (ECM) of these tissues. Enzymatic tissue dissociation is superior to mechanical tissue dissociation as it allows a higher yield of leukocytes with less shear-force-associated damage35. Consequently, mechanical tissue dissociation requires pooling tissues36, which may increase the variability and heterogeneity of the samples. Yet, mechanical dissociation may be the choice when the antigen of interest can be altered by enzymatic dissociation or when the functionality of the cells of interest need to be preserved (e.g., cytotoxicity of NK cells)35.
The use of proteolysis with specific enzymes to degrade the ECM eliminates the low yields observed with mechanical dissociation. Several studies have reported the use of trypsin32, collagenase37, DNase31, dispase38, and commercial cocktails of various enzymes32,39. However, the nature and concentration of different enzymes and the duration of digestion must be meticulously defined and validated in order to ensure maintenance of the integrity of the cell surface antigenic epitopes required for immunophenotyping. The various surface structures are differentially susceptible to destruction by different enzymes, with some enzymes, such as trypsin, being notorious for stripping leukocyte surface epitopes recognized by many monoclonal antibodies.
Introduced herein is a method using a proteolytic and collagenolytic enzymatic cocktail, called Accutase. This enzymatic solution is gentle enough while still efficient in dissociating murine tissues at the maternal-fetal interface, and does not require the addition of other dissociating reagents or serum to terminate the dissociation reaction. Moreover, it is ready to use as supplied and, although the time of dissociation needs to be validated, it is more robust than the above-noted enzymes40,41.
The utilization of a combination of both types of tissue disaggregation improves the quality and amount of cells obtained; thus, several studies have implemented the combined use of mechanical and enzymatic dissociation with satisfactory results31,32,37. The protocol described herein was established and validated in our laboratory; it uses a combination of a gentle mechanical dissociation followed by a robust enzymatic disaggregation. This protocol allows the isolation and further study of the infiltrating leukocytes in murine tissues at the maternal-fetal interface (uterus, decidua, and placenta). The following protocol also maintains the integrity of cell surface markers and yields enough viable cells for downstream applications as demonstrated by flow cytometric analysis. Finally, this protocol maintains the consistency of cell preparation for the analysis and comparison of different murine tissues composing the maternal-fetal interface.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Magentic Cell Separation</td>
		</tr>
		<tr>
			<td>MS Columns</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Separator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>30&#956;m pre separation filters</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Multistand</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>15mL safe lock conical tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MACS Buffer</td>
			<td></td>
			<td></td>
			<td>(0.5% bovine serum albumin, 2mM EDTA and 1X PBS)</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagents</td>
		</tr>
		<tr>
			<td>Anti-mouse CD16/CD32</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-mouse extracellular antibodies</td>
			<td></td>
			<td></td>
			<td>(Table 1)</td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td></td>
			<td></td>
			<td>(BSA)</td>
		</tr>
		<tr>
			<td>LIVE/DEAD viability dye</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fixation buffer solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>FACS Buffer</td>
			<td></td>
			<td></td>
			<td>(1% bovine serum albumin, 0.5% sodium azide, and 1X PBS ph 7.2)</td>
		</tr>
		<tr>
			<td>Trypan Blue Solution 0.4%</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Additional Instruments</td>
		</tr>
		<tr>
			<td>Incubator with shaker</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Flow cytometer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum system</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cell counter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation of leukocytes from the murine tissues at the maternal-fetal interface

Immune tolerance in pregnancy requires that the immune system of the mother undergoes distinctive changes in order to accept and nurture the developing fetus. This tolerance is initiated during coitus, established during fecundation and implantation, and maintained throughout pregnancy. Active cellular and molecular mediators of maternal-fetal tolerance are enriched at the site of contact between fetal and maternal tissues, known as the maternal-fetal interface, which includes the placenta and the uterine and decidual tissues. This interface is comprised of stromal cells and infiltrating leukocytes, and their abundance and phenotypic characteristics change over the course of pregnancy. Infiltrating leukocytes at the maternal-fetal interface include neutrophils, macrophages, dendritic cells, mast cells, T cells, B cells, NK cells, and NKT cells that together create the local micro-environment that sustains pregnancy. An imbalance among these cells or any inappropriate alteration in their phenotypes is considered a mechanism of disease in pregnancy. Therefore, the study of leukocytes that infiltrate the maternal-fetal interface is essential in order to elucidate the immune mechanisms that lead to pregnancy-related complications. Described herein is a protocol that uses a combination of gentle mechanical dissociation followed by a robust enzymatic disaggregation with a proteolytic and collagenolytic enzymatic cocktail to isolate the infiltrating leukocytes from the murine tissues at the maternal-fetal interface. This protocol allows for the isolation of high numbers of viable leukocytes (&#62;70%) with sufficiently conserved antigenic and functional properties. Isolated leukocytes can then be analyzed by several techniques, including immunophenotyping, cell sorting, imaging, immunoblotting, mRNA expression, cell culture, and in vitro functional assays such as mixed leukocyte reactions, proliferation, or cytotoxicity assays.

The dissection of murine tissues from the maternal-fetal interface is shown in Figure 1; this procedure includes opening the peritoneal cavity (Figure 1A,B), uterine horns (Figure 1C)&#160;including the implantation sites (Figure 1D), and the collection of the uterine tissues (Figure 1E), placenta (Figure 1F), and decidual tissues (Figure 1G) at 16.5 dpc. Figure 2 shows the morphology of...

Watch this Scientific Journal Video about Isolation of Leukocytes from the Murine Tissues at the Maternal-Fetal Interface at JoVE.com

Isolation of Leukocytes from the Murine Tissues at the Maternal-Fetal Interface

Described herein is a protocol to isolate and analyze the infiltrating leukocytes of tissues at the maternal-fetal interface (uterus, decidua, and placenta) of mice. This protocol maintains the integrity of most cell surface markers and yields enough viable cells for downstream applications including flow cytometry analysis.

Immune tolerance in pregnancy requires that the immune system of the mother undergoes distinctive changes in order to accept and nurture the developing ...

isolation-leukocytes-from-murine-tissues-at-maternal-fetal

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Immunology and Infection

17.1K Views.  Wayne State University School of Medicine. Described herein is a protocol to isolate and analyze the infiltrating leukocytes of tissues at the maternal-fetal interface (uterus, decidua, and placenta) of mice. This protocol maintains the integrity of most cell surface markers and yields enough viable cells for downstream applications including flow cytometry analysis. 

Video: Isolation of Leukocytes from the Murine Tissues at the Maternal-Fetal Interface

Isolation of Brain-infiltrating Leukocytes

We describe a method for preparing brain infiltrating leukocytes (BILs) from mice. We demonstrate how to infect mice with Theiler's murine encephalomyelitis virus (TMEV) via a rapid intracranial injection technique and how to purify a leukocyte-enriched population of infiltrating cells from whole brain. Briefly, mice are anesthetized with isoflurane in a closed chamber and are free-hand injected with a Hamilton syringe into the frontal cortex. Mice are then killed at various times after infection by isoflurane overdose and whole brains are extracted and homogenized in RPMI with a Tenbroeck tissue grinder. Brain homogenates are centrifuged through a continuous 30% Percoll gradient to remove the myelin and other cell debris. The cell suspension is then strained at 40 &mu;m, washed and centrifuged on a discontinuous Ficoll-Paque Plus gradient to select and purify the leukocytes. The leukocytes are then washed and resuspended in appropriate buffers for immunophenotyping by flow cytometry. Flow cytometry reveals a population of innate immune cells at the early stages of infection in C57BL/6 mice. At 24 hours post infection, multiple subsets of immune cells are present in the BILs, with an enriched population of Gr1+, CD11b+ and F4/80+cells. Therefore, this method is useful in characterizing the immune response to acute infection in the brain.

A rapid method to obtain infiltrating leukocytes from the murine brain is described. This method utilizes a continuous Percoll gradient and discontinuous Ficoll gradient to select and purify the leukocyte-enriched layer. Isolated leukocytes may then be characterized by flow cytometric measurements.

We describe a method for preparing brain infiltrating leukocytes (BILs) from mice. We demonstrate how to infect mice with Theiler's murine ...

RNA Isolation of Pseudomonas aeruginosa Colonizing the Murine Gastrointestinal Tract

Pseudomonas aeruginosa (PA) infections result in significant morbidity and mortality in hosts with compromised immune systems, such as patients with leukemia, severe burn wounds, or organ transplants1. In patients at high-risk for developing PA bloodstream infections, the gastrointestinal (GI) tract is the main reservoir for colonization2, but the mechanisms by which PA transitions from an asymptomatic colonizing microbe to an invasive, and often deadly, pathogen are unclear. Previously, we performed in vivo transcription profiling experiments by recovering PA mRNA from bacterial cells residing in the cecums of colonized mice 3 in order to identify changes in bacterial gene expression during alterations to the host&#x2019;s immune status.
As with any transcription profiling experiment, the rate-limiting step is in the isolation of sufficient amounts of high quality mRNA. Given the abundance of enzymes, debris, food residues, and particulate matter in the GI tract, the challenge of RNA isolation is daunting. Here, we present a method for reliable and reproducible isolation of bacterial RNA recovered from the murine GI tract. This method utilizes a well-established murine model of PA GI colonization and neutropenia-induced dissemination4. Once GI colonization with PA is confirmed, mice are euthanized and cecal contents are recovered and flash frozen. RNA is then extracted using a combination of mechanical disruption, boiling, phenol/chloroform extractions, DNase treatment, and affinity chromatography. Quantity and purity are confirmed by spectrophotometry (Nanodrop Technologies) and bioanalyzer (Agilent Technologies) (Fig 1). This method of GI microbial RNA isolation can easily be adapted to other bacteria and fungi as well.

RNA Isolation of Pseudomonas aeruginosa Colonizing the Murine Gastrointestinal Tract

A reliable method for the RNA isolation of Pseudomonas aeruginosa recovered from murine cecums is described. The RNA recovered is of sufficient quantity and quality for subsequent qPCR, transcription profiling, and RNA Seq experiments. This technique can be adapted for RNA isolation of other intestinal microbes.

Pseudomonas aeruginosa (PA) infections result in significant morbidity and mortality in hosts with compromised immune systems, such as patients with ...

Isolation and Analysis of Brain-sequestered Leukocytes from Plasmodium berghei ANKA-infected Mice

We describe a method for isolation and characterization of adherent inflammatory cells from brain blood vessels of P. berghei ANKA-infected mice. Infection of susceptible mouse-strains with this parasite strain results in the induction of experimental cerebral malaria, a neurologic syndrome that recapitulates certain important aspects of Plasmodium falciparum-mediated severe malaria in humans 1,2 . Mature forms of blood-stage malaria express parasitic proteins on the surface of the infected erythrocyte, which allows them to bind to vascular endothelial cells. This process induces obstructions in blood flow, resulting in hypoxia and haemorrhages 3 and also stimulates the recruitment of inflammatory leukocytes to the site of parasite sequestration.
Unlike other infections, i.e neutrotopic viruses4-6, both malaria-parasitized red blood cells (pRBC) as well as associated inflammatory leukocytes remain sequestered within blood vessels rather than infiltrating the brain parenchyma. Thus to avoid contamination of sequestered leukocytes with non-inflammatory circulating cells, extensive intracardial perfusion of infected-mice prior to organ extraction and tissue processing is required in this procedure to remove the blood compartment. After perfusion, brains are harvested and dissected in small pieces. The tissue structure is further disrupted by enzymatic treatment with Collagenase D and DNAse I. The resulting brain homogenate is then centrifuged on a Percoll gradient that allows separation of brain-sequestered leukocytes (BSL) from myelin and other tissue debris. Isolated cells are then washed, counted using a hemocytometer and stained with fluorescent antibodies for subsequent analysis by flow cytometry.
This procedure allows comprehensive phenotypic characterization of inflammatory leukocytes migrating to the brain in response to various stimuli, including stroke as well as viral or parasitic infections. The method also provides a useful tool for assessment of novel anti-inflammatory treatments in pre-clinical animal models.

Isolation and Analysis of Brain-sequestered Leukocytes from Plasmodium berghei ANKA-infected Mice

A method for isolation of adherent inflammatory leukocytes from brain blood vessels of Plasmodium berghei ANKA-infected mice is described. The method allows quantification as well as phenotypic characterization of isolated leukocytes after staining with fluorescent antibodies and subsequent analysis by flow cytometry.

We describe a method for isolation and  characterization of adherent inflammatory cells from brain blood vessels of P. berghei ANKA-infected mice. ...

Bioenergetics and the Oxidative Burst: Protocols for the Isolation and Evaluation of Human Leukocytes and Platelets

Mitochondrial dysfunction is known to play a significant role in a number of pathological conditions such as atherosclerosis, diabetes, septic shock, and neurodegenerative diseases but assessing changes in bioenergetic function in patients is challenging.&#160;Although diseases such as diabetes or atherosclerosis present clinically with specific organ impairment, the systemic components of the pathology, such as hyperglycemia or inflammation, can alter bioenergetic function in circulating leukocytes or platelets. This concept has been recognized for some time but its widespread application has been constrained by the large number of primary cells needed for bioenergetic analysis.&#160;This technical limitation has been overcome by combining the specificity of the magnetic bead isolation techniques, cell adhesion techniques, which allow cells to be attached without activation to microplates, and the sensitivity of new technologies designed for high throughput microplate respirometry.&#160;An example of this equipment is the extracellular flux analyzer. Such instrumentation typically uses oxygen and pH sensitive probes to measure rates of change in these parameters in adherent cells, which can then be related to metabolism. Here we detail the methods for the isolation and plating of monocytes, lymphocytes, neutrophils and platelets, without activation, from human blood and the analysis of mitochondrial bioenergetic function in these cells. In addition, we demonstrate how the oxidative burst in monocytes and neutrophils can also be measured in the same samples. Since these methods use only 8-20 ml human blood they have potential for monitoring reactive oxygen species generation and bioenergetics in a clinical setting.

Blood leukocytes and platelets can be used as a marker of overall bioenergetic health of an individual and so have the potential to monitor pathological processes and the impact of treatments. Here&#160;we describe a method to isolate and measure mitochondrial function and the oxidative burst in these cells.

Mitochondrial dysfunction is known to play a significant role in a number of pathological conditions such as atherosclerosis, diabetes, septic shock, and ...

Isolation of Human Monocytes by Double Gradient Centrifugation and Their Differentiation to Macrophages in Teflon-coated Cell Culture Bags

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to understand the underlying mechanisms of these diseases. We therefore present a straightforward protocol for the isolation of human monocytes from buffy coats, followed by a differentiation procedure which results in high macrophage yields. The technique relies mostly on commonly available lab equipment and thus provides a cost and time effective way to obtain large quantities of human macrophages. Briefly, buffy coats from healthy blood donors are subjected to a double density gradient centrifugation to harvest monocytes from the peripheral blood. These monocytes are then cultured in fluorinated ethylene propylene (FEP) Teflon-coated cell culture bags in the presence of macrophage colony-stimulating factor (M-CSF). The differentiated macrophages can be easily harvested and used for subsequent studies and functional assays. Important methods for quality control and validation of the isolation and differentiation steps will be highlighted within the protocol. In summary, the protocol described here enables scientists to routinely and reproducibly isolate human macrophages without the need for cost intensive tools. Furthermore, disease models can be studied in a syngeneic human system circumventing the use of murine macrophages.

We present a simple and efficient protocol for the generation of human macrophages. Buffy coats are processed by double density gradient centrifugation and isolated monocytes are then differentiated to macrophages in Teflon-coated cell culture bags. This maximizes macrophage yields and facilitates cell harvesting for subsequent experiments.

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to ...

Analyzing the Effects of Stromal Cells on the Recruitment of Leukocytes from Flow

Stromal cells regulate the recruitment of circulating leukocytes during inflammation through cross-talk with neighboring endothelial cells. Here we describe two in vitro &#8220;vascular&#8221; models for studying the recruitment of circulating neutrophils from flow by inflamed endothelial cells. A major advantage of these models is the ability to analyze each step in the leukocyte adhesion cascade in order, as would occur in vivo. We also describe how both models can be adapted to study the role of stromal cells, in this case mesenchymal stem cells (MSC), in regulating leukocyte recruitment.
Primary endothelial cells were cultured alone or together with human MSC in direct contact on Ibidi microslides or on opposite sides of a Transwell filter for 24 hr. Cultures were stimulated with tumor necrosis factor alpha (TNF&#945;) for 4 hr and incorporated into a flow-based adhesion assay. A bolus of neutrophils was perfused over the endothelium for 4 min. The capture of flowing neutrophils and their interactions with the endothelium was visualized by phase-contrast microscopy.
In both models, cytokine-stimulation increased endothelial recruitment of flowing neutrophils in a dose-dependent manner. Analysis of the behavior of recruited neutrophils showed a dose-dependent decrease in rolling and a dose-dependent increase in transmigration through the endothelium. In co-culture, MSC suppressed neutrophil adhesion to TNF&#945;-stimulated endothelium.
Our flow based-adhesion models mimic the initial phases of leukocyte recruitment from the circulation. In addition to leukocytes, they can be used to examine the recruitment of other cell types, such as therapeutically administered MSC or circulating tumor cells. Our multi-layered co-culture models have shown that MSC communicate with endothelium to modify their response to pro-inflammatory cytokines, altering the recruitment of neutrophils. Further research using such models is required to fully understand how stromal cells from different tissues and conditions (inflammatory disorders or cancer) influence the recruitment of leukocytes during inflammation.

The ability of inflamed endothelium to recruit leukocytes from flow is regulated by mesenchymal stromal cells. We describe two in vitro models incorporating primary human cells that can be used to assess neutrophil recruitment from flow and examine the role that mesenchymal stromal cells play in regulating this process.

Stromal cells regulate the recruitment of circulating leukocytes during inflammation through cross-talk with neighboring endothelial cells. Here we ...

Isolation and Transplantation of Different Aged Murine Thymic Grafts.

The mechanisms that regulate the efficacy of thymic selection remain ill-defined. The method presented here allows in vivo analyses of the development and selection of T cells specific for self and foreign antigens. The approach entails implantation of thymic grafts derived from various aged mice into immunodeficient scid recipients. Over a relatively short period of time the recipients are fully reconstituted with T cells derived from the implanted thymus graft. Only thymocytes seeding the thymus at the time of isolation undergo selection and develop into mature T cells. As such, changes in the nature and specificity of the engrafted T cells as a function of age-dependent thymic events can be assessed. Although technical expertise is required for successful thymic transplantation, this method provides a unique strategy to study in vivo a wide range of pathologies that are due to or a result of aberrant thymic function and/or homeostasis.

This report provides a detailed description of transplanting murine thymi from different aged donor mice under the kidney capsule of immunodeficient mouse recipients. The goal of this approach is to model T cell development and thymic selection events in vivo.

The mechanisms that regulate the efficacy of thymic selection remain ill-defined. The method presented here allows in vivo analyses of the development and ...

Isolation of Leukocytes from the Human Maternal-fetal Interface

Pregnancy is characterized by the infiltration of leukocytes in the reproductive tissues and at the maternal-fetal interface (decidua basalis and decidua parietalis). This interface is the anatomical site of contact between maternal and fetal tissues; therefore, it is an immunological site of action during pregnancy. Infiltrating leukocytes at the maternal-fetal interface play a central role in implantation, pregnancy maintenance, and timing of delivery. Therefore, phenotypic and functional characterizations of these leukocytes will provide insight into the mechanisms that lead to&#160;pregnancy disorders. Several protocols have been described in order to isolate infiltrating leukocytes from the decidua basalis and decidua parietalis; however, the lack of consistency in the reagents, enzymes, and times of incubation makes it difficult to compare these results. Described herein is&#160;a novel approach that combines the use of gentle mechanical and enzymatic dissociation techniques to preserve the viability and integrity of extracellular and intracellular markers in leukocytes isolated from the human tissues at the maternal-fetal interface. Aside from immunophenotyping, cell culture, and cell sorting, the future applications of this protocol are numerous and varied. Following this protocol, the isolated leukocytes can be used to determine DNA methylation, expression of target genes, in vitro leukocyte functionality (i.e., phagocytosis, cytotoxicity, T-cell proliferation, and plasticity, etc.), and the production of reactive oxygen species at the maternal-fetal interface. Additionally, using the described protocol, this laboratory has been able to describe new and rare leukocytes at the maternal-fetal interface.

Described herein is a protocol to isolate and further study the infiltrating leukocytes of the decidua basalis and decidua parietalis - the human maternal-fetal interface. This protocol maintains the integrity of cell surface markers and yields enough viable cells for downstream applications as proven by flow cytometry analysis.

Pregnancy is characterized by the infiltration of leukocytes in the reproductive tissues and at the maternal-fetal interface (decidua basalis and decidua ...

Isolation of Infiltrating Leukocytes from Mouse Skin Using Enzymatic Digest and Gradient Separation

Dissociating murine skin into a single cell suspension is essential for downstream cellular analysis such as the characterization of infiltrating immune cells in rodent models of skin inflammation. Here, we describe a protocol for the digestion of mouse skin in a nutrient-rich solution with collagenase D, followed by separation of hematopoietic cells using a discontinuous density gradient. Cells thus obtained can be used for in vitro studies, in vivo transfer, and other downstream cellular and molecular analyses including flow cytometry. This protocol is an effective and economical alternative to expensive mechanical dissociators, specialized separation columns, and harsher trypsin- and dispase-based digestion methods, which may compromise cellular viability or density of surface proteins relevant for phenotypic characterization or cellular function. As shown here in our representative data, this protocol produced highly viable cells, contained specific immune cell subsets, and had no effect on integrity of common surface marker proteins used in flow cytometric analysis.

This protocol describes enzymatic digestion of mouse skin in nutrient-rich medium followed by gradient separation to isolate leukocytes. Cells thus derived can be used for diverse downstream applications. This is an effective, economical, and improved alternative to tissue dissociation machines and harsher trypsin and dispase-based tissue digestion protocols.

Dissociating murine skin into a single cell suspension is essential for downstream cellular analysis such as the characterization of infiltrating immune ...

Human Placental and Decidual Organ Cultures to Study Infections at the Maternal-fetal Interface

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is imperative to design experiments using human cells and tissues. An advantage of organ culture is maintenance of three-dimensional (3D) structural organization and extracellular matrix. The goal of the method described here is successful establishment of ex vivo human gestational tissue organ cultures and their healthy culture maintenance for 72-96 hr. The protocol details the immediate processing of research-consented, placental and decidual specimens fresh from the operating suite. These are abundant specimens that would otherwise be discarded. Detailed instructions on the sterile collection of these samples, including morphologic details on how to select appropriate tissues to establish 3D organ cultures, is provided. Placental villous and decidual tissues are microdissected into 2-3 mm3 pieces and placed separately on matrix-lined transwell filters and cultured for several days. Villous and decidual organ cultures are well suited for the study of human host-pathogen interaction. As compared to other model organisms, these human cultures are particularly advantageous to examine mechanism of infection for pathogens that demonstrate variable patterns of host specificity. As an example, we demonstrate infection of placental and decidual organ cultures with the clinically relevant, facultative intracellular bacterial pathogen Listeria monocytogenes.

A simple method to establish primary human placental (villous) and decidual organ cultures is described. Villous and decidual organ cultures are invaluable tools for studying pathogenesis at the human maternal-fetal interface. Infection with the facultative intracellular bacterium Listeria monocytogenes is demonstrated.

The placenta shows a large degree of interspecies anatomic variability. To best understand biology and pathophysiology of the human placenta, it is ...