The authors wish to thank Monique Freund, Catherine Ziessel and Ketty for technical assistance. This work was supported by ARMESA (Association de Recherche et D&#233;veloppement en M&#233;decine et Sant&#233; Publique), and by Grant ANR-17-CE14-0001-01 to Henri.de la.Salle.

Protocols involving animals were performed in accordance with the CREMEAS Committee on the Ethics of Animal Experiments of the University of Strasbourg (Comit&#233; R&#233;gional d&#39;Ethique en Mati&#232;re d&#39;Exp&#233;rimentation Animale Strasbourg. Permit Number: E67-482-10).
1. Mouse bone collection
<ol>
	<li>Sacrifice the animal in compliance with the institutional guidelines. 
	NOTE: The data presented in this manuscript were obtained from C57Bl/6 mice of 8 to 12 weeks old. The number of cells obtained, and the frequency of cited populations may vary with age and mouse strain.</li>
	<li>Spray the body with 70% ethanol.</li>
	<li>Using scissors, make a 0.5-1 cm incision of the skin perpendicular to the spine and tear the skin around the whole body. Pull down the skin from the lower body and remove the skin.</li>
	<li>Place the animal on the dissection pad, face down. Locate the pelvic bones by sliding your fingers along the exposed spine from the top to the bottom. To locate the iliac crest, identify the small bump in the lumbar region near the hindlimbs (the anterosuperior region of the pelvic bone). Figure 1A,B presents a schematic representation of the mouse anatomy.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62498/62498fig01.jpg" /> 
Figure 1: Mouse anatomy. (A) Mouse X-Ray showing the hindlimb bones. Note the space between the pelvic bone and the spine (yellow arrow), where the scissors must be inserted to properly separate the hindlimbs from the body of the mouse (yellow dotted line). (B) Schematic representation of the bone marrow-rich bones of interest. The pelvic bones are depicted in red, the femurs in purple, and the tibias in green. (C) Schematic representation of the mouse pelvic bone. The ilium corresponds to the marrow-rich part of the pelvic bone and is highlighted in red. <a href="https://www.jove.com/files/ftp_upload/62498/62498fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="5">
	<li>Place the scissors parallel to the spine against the vertebrae and close to the iliac crest bump. Proceed to cut the muscles along the side of the spine above the pelvic bone by sliding the scissors along the vertebrae all the way down to the tail. 
	NOTE: This first section of muscles can also be performed using a scalpel blade.</li>
	<li>Place the scissors parallel to the spine and proceed to cut between the vertebrae and the iliac crest, as indicated by the yellow dotted line on Figure 1A. Make sure to remain as close to the vertebrae as possible. Cut the remaining muscles to detach the limb from the body. 
	NOTE: There should be little to no resistance.</li>
	<li>Repeat on the other side to detach the second limb.</li>
	<li>Transfer the limbs on a clean surface and discard the rest of the body in compliance with the institutional guidelines.</li>
	<li>Expose the pelvic, femoral, and tibial bones by removing as much surrounding tissue as possible with the forceps and the scalpels.</li>
	<li>Proceed to carefully dislocate the femoral head from the pelvic bone by holding the distal end of the femur with the forceps while gently slicing the muscles around the articulation with the scalpels. Wiggle the bones to facilitate the dislocation.</li>
	<li>Scrape off the remaining muscle from the pelvic bone and cut with a scalpel in the middle of the cavity that did hold the femur head. The ilium is kept as it is rich in hematopoietic progenitors while the triangular very thin side of the bone is discarded, as shown in Figure 1C.</li>
	<li>Remove the residual tissues around the ilium with the scalpel and place the cleaned bone in sterile PBS supplemented with 2% Newborn Calf Serum (PBS-2%NBCS).</li>
	<li>Using scissors, cut off the foot from the leg at the ankle.</li>
	<li>Hold the lower part of the tibia with the forceps and scrape the muscle up toward the knee. Discard the fibulae and cut across the tibial plateau with the scalpel. Place the tibia in sterile PBS-2%NBCS.</li>
	<li>Remove the residual tissues around the femur with scalpels.</li>
	<li>Hold the upper side of the femur with forceps; place the scalpel blade at the base of the kneecap. Apply a force toward the kneecap parallel to the femur until the detachment of the kneecap. Place the femur in sterile PBS-2%NBCS. Removal of the kneecap provides a clean access for inserting the needle for marrow flushing.</li>
</ol>
2. Magnetic depletion of lineage positive cells
<ol>
	<li>In a laminar flow cabinet, transfer the bones in a sterile Petri dish filled with sterile PBS-2%NBCS.</li>
	<li>With a scalpel cut off the head of the femurs.</li>
	<li>Fill a 1 mL syringe with sterile PBS-2%NBCS and attach a 21 G needle to the outlet.</li>
	<li>Fill a 5 mL polypropylene tube with 2 mL of sterile PBS-2%NBCS.</li>
	<li>Hold the femur with the forceps; gently insert the needle in the groove left after the kneecap removal. Apply rotation to the needle while inserting to avoid plugging of the needle. Ensure that the needle is completely inserted into the bone up to the bevel.</li>
	<li>Transfer the bone with the needle into the tube containing 2 mL of PBS-2%NBCS. Dispense and aspirate the PBS-2%NBCS from the syringe until the bone is clear.</li>
	<li>Remove the needle from the femur and insert it in the hole at opposite side where the femur head was. Dispense and aspirate the buffer again and discard the bone.</li>
	<li>For the iliac crest and tibia, hold the bone with the forceps; gently insert the needle in the open side. Apply rotation to the needle while inserting to avoid plugging of the needle. Ensure that the needle is completely inserted into the bone up to the bevel. Transfer the bone with the needle into the tube containing 2 mL of PBS-2%NBCS. Dispense and aspirate the PBS-2%NBCS from the syringe until the bone is clear. Discard the bones. 
	NOTE: Bones from up to three mice can be flushed into the same tube. Pool the cell suspensions.</li>
	<li>Pass the pooled cell suspension through a 40 &#181;m cell strainer cap placed onto a sterile 5 mL polystyrene tube.</li>
	<li>Proceed to count the cells. 
	NOTE: Cell count can be performed with any hemocytometer, using Trypan Blue for viability assessment, or with any automated cell counter. One mouse typically yields 105 &#177; 7 x 106 cells.</li>
	<li>Take aside 100 &#181;L of the cell suspension as Total Bone Marrow, add 500 &#181;L of PBS-2%NBCS and save it on ice for the staining procedure.</li>
	<li>Pellet the filtered suspension by centrifugation at 400 x g for 5 min at 4 &#176;C and discard the supernatant. 
	NOTE: Red blood cells can be lysed by resuspending the pellet in freshly prepared Lysis Solution (1/10th in dH2O). Incubate for 5 min until the suspension becomes clear and bright red and add 10 volumes of sterile PBS. Proceed to wash the cells in PBS-2%NBCS by centrifugation at 400 x g for 5 mins at 4 &#176;C. Be careful when removing the supernatant as the cell pellet is very loose. Perform a second wash with PBS-2%NBCS by centrifugation at 400 x g for 5 min at 4 &#176;C and proceed to step 2.13.</li>
	<li>Resuspend the cell pellet in freshly prepared primary antibody cocktail with a ratio of 100 &#181;L per 1 x 107 cells. Incubate on ice for 30-45 min.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Antibody</td>
			<td>Dilution</td>
		</tr>
		<tr>
			<td>Gr-1-biotin</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>B220-biotin</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>Mac-1-biotin</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>CD3-biotin</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>CD4-biotin</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>CD5-biotin</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>CD8-biotin</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>TER119-biotin</td>
			<td>1:1000</td>
		</tr>
		<tr>
			<td>CD127-biotin</td>
			<td>1:500</td>
		</tr>
	</tbody>
</table>
Table 1.
<ol start="14">
	<li>Take aside 10 &#181;L of the cell suspension into a sterile 5 mL polystyrene tube labeled Lin-Pos Fraction. Add 90 &#181;L of PBS-2%NBCS and save it on ice for the staining procedure.</li>
	<li>Proceed to wash the cells twice with sterile PBS-2%NBCS by centrifugation at 400 x g for 5 min at 4 &#176;C. Make sure to do the last wash in a sterile 5 mL polypropylene tube.</li>
	<li>During the washing steps, prepare the beads for the magnetic depletion.
	<ol>
		<li>Resuspend the beads in the vial by thoroughly vortexing for 30 s.</li>
		<li>Transfer a volume of beads corresponding to two beads per target cell into a 5 mL polypropylene tube.</li>
		<li>Wash the beads twice with PBS-2%NBCS by placing the tube on the magnet and removing the washing buffer using a sterile glass Pasteur pipette.</li>
		<li>Resuspend the beads in 500 &#181;L of sterile PBS-2NBCS%.</li>
	</ol>
	</li>
	<li>Resuspend the pellet of labeled cells in 250 &#181;L of beads and mix gently for 5 min on ice. Add 2 mL of sterile PBS-2%NBCS and mix gently. Do not shake the tube.</li>
	<li>Place the tube onto the magnet for 2 min.</li>
	<li>Proceed to collect the non-magnetic fraction with a sterile glass Pasteur pipette and add it onto the remaining 250 &#181;L of magnetic beads. Seal the tube with parafilm.</li>
	<li>Place the tube on a tube roller for 20 min at 4 &#176;C.</li>
	<li>Add 2 mL of sterile PBS-2%NBCS and mix gently. Do not shake the tube.</li>
	<li>Place the tube in the magnet for 2 min.</li>
	<li>Proceed to collect the non-magnetic fraction into a sterile 5 mL polypropylene tube labeled Lin-Neg Fraction with a sterile glass Pasteur pipette.</li>
	<li>Pellet the cells by centrifugation at 400 x g for 5 min at 4 &#176;C and remove the supernatant.</li>
	<li>Resuspend the non-magnetic cells in 500 &#181;L of sterile PBS-2%NBCS.</li>
	<li>Proceed to count the cells. 
	&#8203;NOTE: One mouse typically yields 3.9 &#177; 1.1 x 106 cells. Typical lineage staining pre- and post- depletion are presented in Figure 2B.</li>
</ol>
3. Cell sorting of megakaryocyte progenitors by flow cytometry
<ol>
	<li>Take the tubes labeled Total Bone Marrow, Lin-Pos Fraction, and Lin-Neg Fraction.</li>
	<li>Proceed to split the content of the tube Total Bone Marrow equally into six sterile 5 mL polystyrene tubes. Label the tubes with the numbers 1-6.</li>
	<li>Proceed to label the tube Lin-Pos Fraction with the number 7.</li>
	<li>Proceed to split the content of the tube Lin-Neg Fraction as follows.
	<ol>
		<li>Transfer 50 &#181;L into a sterile 5 mL polystyrene tube containing 250 &#181;L of sterile PBS-2%NBCS. Then, split its content equally into 3 sterile 5 mL polystyrene tubes. Label these tubes with the numbers 8-10.</li>
		<li>The remaining 450 &#181;L of Lin-Neg Fraction cell suspension corresponds to the tube with the number 11.</li>
	</ol>
	</li>
	<li>Add the antibodies to the tubes as described in Table 2.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Tube</td>
			<td>Label</td>
			<td>Antibody cocktail</td>
		</tr>
		<tr>
			<td colspan="3">Total Bone Marrow</td>
		</tr>
		<tr>
			<td>1</td>
			<td>Unstained control</td>
			<td></td>
		</tr>
		<tr>
			<td>2</td>
			<td>Single stained control</td>
			<td>CD45-FITC (1/200)</td>
		</tr>
		<tr>
			<td>3</td>
			<td>Single stained control</td>
			<td>CD45-PE (1/200)</td>
		</tr>
		<tr>
			<td>4</td>
			<td>Single stained control</td>
			<td>TER119-APC (1/200)</td>
		</tr>
		<tr>
			<td>5</td>
			<td>Single stained control</td>
			<td>CD45-PECy7 (1/200)</td>
		</tr>
		<tr>
			<td>6</td>
			<td>Single stained control</td>
			<td>CD45-APC-Cy7 biotin (1/200)</td>
		</tr>
		<tr>
			<td colspan="3">Lin-Pos Fraction</td>
		</tr>
		<tr>
			<td>7</td>
			<td>Single stained control</td>
			<td>Single stained control. Streptavidin-APC-Cy7 (1/500)</td>
		</tr>
		<tr>
			<td colspan="3">Lin-Neg Fraction</td>
		</tr>
		<tr>
			<td>8</td>
			<td>FMO FITC control</td>
			<td>&#160;c-kit-APC (1/200) + Sca-1-PE (1/200) + 
			CD16/32-PE (1/200) + CD150-PECy7 (1/200) + Streptavidin-APC-Cy7 (1/500)</td>
		</tr>
		<tr>
			<td>9</td>
			<td>FMO PE control</td>
			<td>CD9-FITC (1/200) + c-kit-APC (1/200) + CD150-PECy7 (1/200) + Streptavidin-APC-Cy7 (1/500)</td>
		</tr>
		<tr>
			<td>10</td>
			<td>FMO PECy7 control</td>
			<td>CD9-FITC (1/200) + c-kit-APC (1/200) + Sca-1-PE (1/200) + 
			CD16/32-PE (1/200) + Streptavidin-APC-Cy7 (1/500)</td>
		</tr>
		<tr>
			<td>11</td>
			<td>Positive tube for sorting</td>
			<td>CD9-FITC (1/200) + c-kit-APC (1/200) + Sca-1-PE (1/200) + 
			CD16/32-PE (1/200) + CD150-PECy7 (1/200) + Streptavidin-APC-Cy7 (1/500)</td>
		</tr>
	</tbody>
</table>
Table 2.
<ol start="6">
	<li>Incubate on ice for 30-45 min in the dark.</li>
	<li>Wash the cells with sterile PBS-2%NBCS by centrifugation at 400 x g for 5 min at 4 &#176;C.</li>
	<li>Resuspend the cell pellets as follows.
	<ol>
		<li>For the tubes 1 to 10, resuspend the pellet in 300 &#181;L of sterile PBS-2%NBCS supplemented with 7AAD (2.5 &#181;g/mL final) (PBS-7AAD). 
		CAUTION: 7AAD is a DNA intercalant and must therefore be handled with appropriate PPE (gloves).</li>
		<li>For tube 11, resuspend the pellet in sterile PBS-7AAD at a maximum concentration of 5 x 106 cells per mL and a minimum volume of 1 mL.</li>
	</ol>
	</li>
	<li>Prepare two polypropylene collection tubes labeled MEP and MKp containing 2 mL of PBS-2%NBCS. 
	NOTE: Alternatively, cells can be collected into culture medium or cell lysis buffer depending on the subsequent application for the sorted cells. The use of polystyrene tubes is not recommended because of possible interference with the charged droplets containing the cells of interest.</li>
	<li>Keep all tubes on ice in the dark.</li>
	<li>Proceed to the cell sorter set-up.
	<ol>
		<li>Use the tubes 1-7 to set-up voltage and compensation, tubes 7-10 to determine the sorting gates for the cell populations of interest and tube 11 for cell sorting.</li>
	</ol>
	</li>
	<li>The first steps of the gating strategy aim to exclude doublets and dead cells from the analysis, as described in Figure 3. Identify single viable cells and display SSC-vs Lin-APC-Cy7 dot plot to confirm the efficiency of the lineage depletion. From the Lin- cells a gate is set to select cells positive for c-kit&#160;and negative or dim for Sca-1 and CD16/32. A CD9 vs CD150 expression dot plot for the selected cells allows the identification of four populations. 
	NOTE: MEP and MKp cells are both positive for CD150. Three levels of expression for CD9 can be defined (neg, dim, and high). MKp express high level of CD9 and MEP express CD9 at an intermediate fluorescence intensity level. MEP population corresponds to Lin- c-Kit+ Sca-1-CD16/32-/dim CD150+ CD9dim and MKp population corresponds to Lin- c-Kit+ Sca-1-CD16/32-/dim CD150+ CD9bright. The discrimination between CD9 high and CD9 dim populations for the CD150 positive cells is set based on the maximum level of CD9 expression in the CD150 negative population. One mouse typically yields 5.3 &#177; 0.6 x 103 MKp and 27.2 &#177; 2.4 x 103 MEP.</li>
</ol>

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The authors declare no competing financial interests.

The method described in this paper allows for the extraction and purification of mouse MEP and MKp. An important parameter in the optimization of the protocol was to obtain sufficient number of cells that would be compatible with most molecular- and cellular-based assays. The general practice of mouse bone collection for hematopoietic cell extraction usually consists in harvesting both the femurs and tibias of each mouse. The pelvic bone, another source of hematopoietic material, is thus often overlooked. The reasons for...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62498">Isolation of Mouse Megakaryocyte Progenitors</a>. A figure was updated.
Figure 2 was updated from:
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62498/62498fig02.jpg" /> 
Figure 2: Magnetic depletion of lineage committed (Lin) cells. (A) Schematic representation of the magnetic depletion protocol. First, unsorted bone marrow cells are labeled with the biotin-conjugated rat anti-mouse antibody cocktail. Cells are then incubated with anti-rat Ig coated magnetic beads and subsequently subjected to the magnetic depletion using a strong magnet. The magnet will retain the labeled magnetic Lin+ fraction against the tube walls, while the unlabeled non-magnetic Lin- negative fraction will be collected in a new tube. (B) Lineage committed cells can be identified using fluorescent conjugated streptavidin. Typical analysis of the lineage expression in cells prior to magnetic depletion (total bone marrow) and after magnetic depletion (Lin- Fraction) N = 21. <a href="https://www.jove.com/files/ftp_upload/62498/62498fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
to:
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62498/62498fig2v2.jpg" /> 
Figure 2: Magnetic depletion of lineage committed (Lin) cells. (A) Schematic representation of the magnetic depletion protocol. First, unsorted bone marrow cells are labeled with the biotin-conjugated rat anti-mouse antibody cocktail. Cells are then incubated with anti-rat Ig coated magnetic beads and subsequently subjected to the magnetic depletion using a strong magnet. The magnet will retain the labeled magnetic Lin+ fraction against the tube walls, while the unlabeled non-magnetic Lin- negative fraction will be collected in a new tube. (B) Lineage committed cells can be identified using fluorescent conjugated streptavidin. Typical analysis of the lineage expression in cells prior to magnetic depletion (total bone marrow) and after magnetic depletion (Lin- Fraction) N = 21. <a href="https://www.jove.com/files/ftp_upload/62498/62498fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62498">Isolation of Mouse Megakaryocyte Progenitors</a>. A figure was updated.

Erratum: Isolation of Mouse Megakaryocyte Progenitors

Blood platelets are produced by megakaryocytes. These large polyploid cells are located in the bone marrow and as for all blood cells they are derived from Hematopoietic Stem Cells (HSC)1. The classical pathway of production of megakaryocytes in the bone marrow originates from HSC and involves the generation of different progenitors that progressively restrict their differentiation potential2. The first progenitor signing the commitment to the megakaryocytic lineage is the Megakaryocyte-Erythrocyte Progenitor (MEP), a bipotent progenitor capable of producing both erythroid cells and megakaryocytes3,4,5. The MEP then produces a unipotent progenitor/precursor (MKp) that will differentiate into a mature megakaryocyte capable of producing platelets. The mechanisms involved in the generation of these progenitors, as well as their differentiation and maturation into megakaryocytes are complex and only partially understood. In addition, the heterogeneity of the MEP population in terms of differentiation potential and the intrinsic commitment level of these cells are still unclear. To decipher these processes, it is essential to obtain (or have access to) purified populations of MEP and MKp for fine molecular and single cell analyses.
Several studies have demonstrated particular combinations of cell surface markers for the identification of progenitors committed to the megakaryocytic lineage in the mouse6,7,8. From these a method was devised allowing the purification of MEP and MKp from mice. This method was optimized to obtain cells in adequate number and quality for a large number of assays. With ethical considerations in mind, and in order to minimize the number of animals involved in the experiments, we elicited to harvest the bone marrow from the femur and tibia, and also from the iliac crest. This bone contains a high frequency and number of hematopoietic progenitors and is most of the time damaged during long bone harvesting. Presented here is a detailed method for the reliable collection of this bone.
The second criteria of optimization is to produce highly purified cell populations. Fluorescent Activated Cell Sorting (FACS) is a method of choice in order to obtain purified populations of cells of interest. However, low yields are reached when the cell population of interest is very rare. Enrichment procedures are thus necessary. In this protocol, a negative selection procedure was opted using magnetic beads.

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isolation of mouse megakaryocyte progenitors

Bone marrow megakaryocytes are large polyploid cells that ensure the production of blood platelets. They arise from hematopoietic stem cells through megakaryopoiesis. The final stages of this process are complex and classically involve the bipotent Megakaryocyte-Erythrocyte Progenitors (MEP) and the unipotent Megakaryocyte Progenitors (MKp). These populations precede the formation of bona fide megakaryocytes and, as such, their isolation and characterization could allow for the robust and unbiased analysis of megakaryocyte formation. This protocol presents in detail the procedure to collect hematopoietic cells from mouse bone marrow, the enrichment of hematopoietic progenitors through magnetic depletion and finally a cell sorting strategy that yield highly purified MEP and MKp populations. First, bone marrow cells are collected from the femur, the tibia, and also the iliac crest, a bone that contains a high number of hematopoietic progenitors. The use of iliac crest bones drastically increases the total cell number obtained per mouse and thus contributes to a more ethical use of animals. A magnetic lineage depletion was optimized using 450 nm magnetic beads allowing a very efficient cell sorting by flow cytometry. Finally, the protocol presents the labeling and gating strategy for the sorting of the two highly purified megakaryocyte progenitor populations: MEP (Lin-Sca-1-c-Kit+CD16/32-CD150+CD9dim) and MKp (Lin- Sca-1-c-Kit+CD16/32-CD150+CD9bright). This technique is easy to implement and provides enough cellular material to perform i) molecular characterization for a deeper knowledge of their identity and biology, ii) in vitro differentiation assays, that will provide a better understanding of the mechanisms of maturation of megakaryocytes, or iii) in vitro models of interaction with their microenvironment.

Phenotypic analysis of the cells identified as MEP and MKp were performed by flow cytometry. Cells were labeled with fluorescence conjugated antibodies to CD41a and CD42c, classical markers of the megakaryocytic and platelet lineages. Both markers were expressed by the cells of the MKp population while these markers are not yet detected at the surface of the cells of the MEP population (Figure 4Ai,4Aii). Polyploidy is a hallmark of megakaryocytes. The DNA content of the sort...

Watch this Scientific Journal Video about Isolation of Mouse Megakaryocyte Progenitors at JoVE.com

Isolation of Mouse Megakaryocyte Progenitors

This method describes the purification by flow cytometry of MEP and MKp from mice femurs, tibias, and pelvic bones.

Bone marrow megakaryocytes are large polyploid cells that ensure the production of blood platelets. They arise from hematopoietic stem cells through ...

isolation-of-mouse-megakaryocyte-progenitors

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6.2K Views.  Université de Strasbourg, INSERM UMR S1255, EFS Grand-EST. This method describes the purification by flow cytometry of MEP and MKp from mice femurs, tibias, and pelvic bones.

Video: Isolation of Mouse Megakaryocyte Progenitors

Mouse Adrenal Chromaffin Cell Isolation

Adrenal medullary chromaffin cell culture systems are extremely useful for the study of excitation-secretion coupling in an in vitro setting.  This protocol illustrates the method used to dissect the adrenals and then isolate the medullary region by stripping away the adrenal cortex.  The digestion of the medulla into single chromaffin cells is then demonstrated.

Adrenal medullary chromaffin cell culture systems are extremely useful for the study of excitation-secretion coupling in an in vitro setting. This protocol illustrates the method used to dissect the adrenals and then isolate the medullary region by stripping away the adrenal cortex. The digestion of the medulla into single chromaffin cells is then demonstrated.

Mysz nadnerczy izolacji komórek chromochłonnych

Adrenal medullary chromaffin cell culture systems are extremely useful for the study of excitation-secretion coupling in an in vitro setting.  This ...

Mouse Dorsal Forebrain Explant Isolation

This video demonstrates the protocol for isolating and culturing explants of the mouse forebrain from embyonic day 12 mice. Procedures for removal of the uterus, embryos from uterus, and dissection of embryos are given. In addition the methodology for transferring these explants onto specialized membranes on which they are cultured is demonstrated. The development of the forebrain can be studied in vitro using this preparations as well as changes in gene expression.

Isolation of Genomic DNA from Mouse Tails

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Purification of Progenitors from Skeletal Muscle

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing in a "satellite cell position" between the myofiber plasma membrane and the laminin-rich basement membrane that ensheaths it, are self-renewing cells that are solely committed to the myogenic lineage1,2. We have recently described a second class of vessel associated progenitors that can generate myofibroblasts and white adipocytes, which responds to damage by efficiently entering proliferation and provides trophic support to myogenic cells during tissue regeneration3,4. One of the most trusted assays to determine the developmental and regenerative potential of a given cell population relies on their isolation and transplantation5-7. To this end we have optimized protocols for their purification by flow cytometry from enzymatically dissociated muscle, which we will outline in this article. The populations obtained using this method will contain either myogenic or fibro/adipogenic colony forming cells: no other cell types are capable of expanding in vitro or surviving in vivo delivery. However, when these populations are used immediately after the sort for molecular analysis (e.g qRT-PCR) one must keep in mind that the freshly sorted subsets may contain other contaminant cells that lack the ability of forming colonies or engrafting recipients.

Method for the enzymatic dissociation, surface labeling and purification by flow cytometry of fibro/adipogenic and myogenic progenitors from murine skeletal muscle.

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing ...

Isolation of Mouse Salivary Gland Stem Cells

Mature salivary glands of both human and mouse origin comprise a minimum of five cell types, each of which facilitates the production and excretion of saliva into the oral cavity. Serous and mucous acinar cells are the protein and mucous producing factories of the gland respectively, and represent the origin of saliva production. Once synthesised, the various enzymatic and other proteinaceous components of saliva are secreted through a series of ductal cells bearing epithelial-type morphology, until the eventual expulsion of the saliva through one major duct into the cavity of the mouth. The composition of saliva is also modified by the ductal cells during this process.
In the manifestation of diseases such as Sj&ouml;gren's syndrome, and in some clinical situations such as radiotherapy treatment for head and neck cancers, saliva production by the glands is dramatically reduced 1,2. The resulting xerostomia, a subjective feeling of dry mouth, affects not only the ability of the patient to swallow and speak, but also encourages the development of dental caries and can be socially debilitating for the sufferer.
The restoration of saliva production in the above-mentioned clinical conditions therefore represents an unmet clinical need, and as such several studies have demonstrated the regenerative capacity of the salivary glands 3-5. Further to the isolation of stem cell-like populations of cells from various tissues within the mouse and human bodies 6-8, we have shown using the described method that stem cells isolated from mouse salivary glands can be used to rescue saliva production in irradiated salivary glands 9,10. This discovery paves the way for the development of stem cell-based therapies for the treatment of xerostomic conditions in humans, and also for the exploration of the salivary gland as a microenvironment containing cells with multipotent self-renewing capabilities.

An optimized protocol for the isolation of stem cells from the mouse salivary gland is described. The method employs enzymatic and mechanical digestion, and permits isolation of salispheres containing cells with characteristics of stem cells.

Mature salivary glands of both human and mouse origin comprise a minimum of five cell types, each of which facilitates the production and excretion of ...

Identification and Analysis of Mouse Erythroid Progenitors using the CD71/TER119 Flow-cytometric Assay

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of red cell formation is regulated by the hormone erythropoietin (Epo), whose synthesis is triggered by tissue hypoxia. A threat to adequate tissue oxygenation results in a rapid increase in Epo, driving an increase in erythropoietic rate, a process known as the erythropoietic stress response. The resulting increase in the number of circulating red cells improves tissue oxygen delivery. An efficient erythropoietic stress response is therefore critical to the survival and recovery from physiological and pathological conditions such as high altitude, anemia, hemorrhage, chemotherapy or stem cell transplantation. 
The mouse is a key model for the study of erythropoiesis and its stress response. Mouse definitive (adult-type) erythropoiesis takes place in the fetal liver between embryonic days 12.5 and 15.5, in the neonatal spleen, and in adult spleen and bone marrow. Classical methods of identifying erythroid progenitors in tissue rely on the ability of these cells to give rise to red cell colonies when plated in Epo-containing semi-solid media. Their erythroid precursor progeny are identified based on morphological criteria. Neither of these classical methods allow access to large numbers of differentiation-stage-specific erythroid cells for molecular study. Here we present a flow-cytometric method of identifying and studying differentiation-stage-specific erythroid progenitors and precursors, directly in the context of freshly isolated mouse tissue. The assay relies on the cell-surface markers CD71, Ter119, and on the flow-cytometric 'forward-scatter' parameter, which is a function of cell size. The CD71/Ter119 assay can be used to study erythroid progenitors during their response to erythropoietic stress in vivo, for example, in anemic mice or mice housed in low oxygen conditions. It may also be used to study erythroid progenitors directly in the tissues of genetically modified adult mice or embryos, in order to assess the specific role of the modified molecular pathway in erythropoiesis.

A flow-cytometric method for identification and molecular analysis of differentiation-stage-specific murine erythroid progenitors and precursors, directly in freshly &#x2013;harvested mouse bone marrow, spleen or fetal liver. The assay relies on cell-surface markers CD71, Ter119, and cell size.

The study of erythropoiesis aims to understand how red cells are formed from earlier hematopoietic and erythroid progenitors. Specifically, the rate of ...

Phenotypic Analysis and Isolation of Murine Hematopoietic Stem Cells and Lineage-committed Progenitors

The bone marrow is the principal site where HSCs and more mature blood cells lineage progenitors reside and differentiate in an adult organism. HSCs constitute a minute cell population of pluripotent cells capable of generating all blood cell lineages for a life-time1. The molecular dissection of HSCs homeostasis in the bone marrow has important implications in hematopoiesis, oncology and regenerative medicine. We describe the labeling protocol with fluorescent antibodies and the electronic gating procedure in flow cytometry to score hematopoietic progenitor subsets and HSCs distribution in individual mice (Fig. 1). In addition, we describe a method to extensively enrich hematopoietic progenitors as well as long-term (LT) and short term (ST) reconstituting HSCs from pooled bone marrow cell suspensions by magnetic enrichment of cells expressing c-Kit. The resulting cell preparation can be used to sort selected subsets for in vitro and in vivo functional studies (Fig. 2).
Both trabecular osteoblasts2,3 and sinusoidal endothelium4 constitute functional niches supporting HSCs in the bone marrow. Several mechanisms in the osteoblastic niche, including a subset of N-cadherin+ osteoblasts3 and interaction of the receptor tyrosine kinase Tie2 expressed in HSCs with its ligand angiopoietin-15 concur in determining HSCs quiescence. "Hibernation" in the bone marrow is crucial to protect HSCs from replication and eventual exhaustion upon excessive cycling activity6. Exogenous stimuli acting on cells of the innate immune system such as Toll-like receptor ligands7 and interferon-&alpha;6 can also induce proliferation and differentiation of HSCs into lineage committed progenitors. Recently, a population of dormant mouse HSCs within the lin- c-Kit+ Sca-1+ CD150+ CD48- CD34- population has been described8. Sorting of cells based on CD34 expression from the hematopoietic progenitors-enriched cell suspension as described here allows the isolation of both quiescent self-renewing LT-HSCs and ST-HSCs9. A similar procedure based on depletion of lineage positive cells and sorting of LT-HSC with CD48 and Flk2 antibodies has been previously described10. In the present report we provide a protocol for the phenotypic characterization and ex vivo cell cycle analysis of hematopoietic progenitors, which can be useful for monitoring hematopoiesis in different physiological and pathological conditions. Moreover, we describe a FACS sorting procedure for HSCs, which can be used to define factors and mechanisms regulating their self-renewal, expansion and differentiation in cell biology and signal transduction assays as well as for transplantation.

A method to analyse the distribution of bone marrow hematopoietic progenitors in flow cytometry as well as to efficiently isolate highly purified hematopoietic stem cells (HSCs) is described. The isolation procedure is essentially based on magnetic enrichment of c-Kit+ cells and cell sorting to purify HSCs for cellular and molecular studies.

The bone marrow is the principal site where HSCs and more mature blood cells lineage progenitors reside and differentiate in an adult organism. HSCs ...

Isolation and Culture of Neonatal Mouse Cardiomyocytes

Cultured neonatal cardiomyocytes have long been used to study myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow for easy investigation and manipulation of biochemical pathways, and their effect on the biomechanical properties of spontaneously beating cardiomyocytes.	
The following 2-day protocol describes the isolation and culture of neonatal mouse cardiomyocytes. We show how to easily dissect hearts from neonates, dissociate the cardiac tissue and enrich cardiomyocytes from the cardiac cell-population. We discuss the usage of different enzyme mixes for cell-dissociation, and their effects on cell-viability. The isolated cardiomyocytes can be subsequently used for a variety of morphological, electrophysiological, biochemical, cell-biological or biomechanical assays. We optimized the protocol for robustness and reproducibility, by using only commercially available solutions and enzyme mixes that show little lot-to-lot variability. We also address common problems associated with the isolation and culture of cardiomyocytes, and offer a variety of options for the optimization of isolation and culture conditions.

Primary mouse cardiomyocyte cultures are one of the pivotal tools for the investigation of myofibrillar organization and function. The following protocol describes the isolation and culture of primary cardiomyocytes from neonatal mouse hearts. The resulting cardiomyocyte cultures may be subsequently used for a variety of biomechanical, biochemical and cell-biological assays.

Cultured neonatal cardiomyocytes have long been used to study  myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow  for easy ...

Isolation and Physiological Analysis of Mouse Cardiomyocytes

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force used to pump blood. Individual cardiomyocytes were first isolated over 40 years ago in order to better study the physiology and structure of heart muscle. Techniques have rapidly improved to include enzymatic digestion via coronary perfusion. More recently, analyzing the contractility and calcium flux of isolated myocytes has provided a vital tool in the cellular and sub-cellular analysis of heart failure. Echocardiography and EKGs provide information about the heart at an organ level only. Cardiomyocyte cell culture systems exist, but cells lack physiologically essential structures such as organized sarcomeres and t-tubules required for myocyte function within the heart. In the protocol presented here, cardiomyocytes are isolated via Langendorff perfusion. The heart is removed from the mouse, mounted via the aorta to a cannula, perfused with digestion enzymes, and cells are introduced to increasing calcium concentrations. Edge and sarcomere detection software is used to analyze contractility, and a calcium binding fluorescent dye is used to visualize calcium transients of electrically paced cardiomyocytes; increasing understanding of the role cellular changes play in heart dysfunction. Traditionally used to test drug effects on cardiomyocytes, we employ this system to compare myocytes from WT mice and mice with a mutation that causes dilated cardiomyopathy. This protocol is unique in its comparison of live cells from mice with known heart function and known genetics. Many experimental conditions are reliably compared, including genetic or environmental manipulation, infection, drug treatment, and more. Beyond physiologic data, isolated cardiomyocytes are easily fixed and stained for cytoskeletal elements. Isolating cardiomyocytes via perfusion is an extremely versatile method, useful in studying cellular changes that accompany or lead to heart failure in a variety of experimental conditions.

Individual cardiomyocytes from wild type and mutant mice can be isolated from the heart in order to study their contractility and calcium transients. This allows characterization of the contribution of cellular dysfunction to heart dysfunction from any cause.

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force ...