Density centrifugation medium, such as Ficoll™ or Lymphoprep™, is widely used for the preparation of mononuclear cell fractions from peripheral blood, cord blood or bone marrow.

Protocol

1. Mix density gradient medium (e.g. Ficoll™, Lymphoprep™) thoroughly before use. Add density gradient medium to an empty tube (see Table 1).
   Note: density gradient medium, blood and medium should remain at room temperature for optimum results.
2. Dilute the blood 2X with PBS + 2% FBS (see Table 1).
3. Layer the blood on top of density gradient medium, being careful to minimize mixing of the blood with the density gradient medium.
4. Centrifuge at room temperature at 400 x g for 30 minutes with the brake off. (G to RPM conversion)
5. Remove and discard the upper plasma layer.
6. Remove and retain the mononuclear cell layer at the plasma-density gradient medium interface.
7. Add PBS + 2% FBS and centrifuge at 300 x g for 8 minutes.
8. Discard the supernatant and resuspend the resultant mononuclear cells in appropriate medium (e.g. PBS + 2% FBS).

Table 1. Recommended Volumes and Tube Sizes

Blood (mL)	PBS + 2% FBS (mL)	Density Gradient Medium (mL)	Tube Size (mL)
1	1	1.5	5
2	2	3	14
3	3	3	14
4	4	4	14
5	5	10	50
10	10	15	50
15	15	15	50

Ficoll™ is a trademark of GE Healthcare Ltd. Lymphoprep™ is a trademark of Nycomed.

Separation Principle
RBC aggregation agents such as HetaSep™ increase the RBC sedimentation rate by increasing the effective size of the cells through formation of aggregates, or rouleaux. Because nucleated cells settle at a lower rate, a compact pellet consisting mainly of RBC is formed rapidly in the presence of HetaSep™, while the nucleated cells remain suspended in the supernatant.

Enriching for nucleated cells from whole blood can speed downstream immunomagnetic cell isolation. Leukocyte-rich plasma can be prepared from peripheral blood samples by sedimentation of red blood cells (RBC) using HetaSep™ by either gravity sedimentation or centrifugation. An interface forms between the RBC fraction and the RBC-depleted (nucleated cell-rich) fraction as the RBCs sediment through the HetaSep™ solution. Approximately 99% RBC depletion is attained if the nucleated cell-rich fraction is removed carefully.

Note: The age of the blood sample impacts how fast and to what extent the RBCs sediment. Accordingly, the interface between the plasma fraction and the RBC fraction may be less distinct in older samples. Haemolysis also makes visualization of the interface less obvious.

Gravity Sedimentation Protocol

1. Based on blood sample volume, select an appropriately sized tube (Table 1).
2. Add 1 part HetaSep™ solution to 5 parts blood. Mix well. If using a blood bag, add HetaSep™ directly to the bag and mix.
3. Allow sample to settle until the plasma:RBC interface is at approximately 50% of the total volume. Placing the tube in a 37°C incubator for this step will increase the sedimentation rate.
4. Harvest the leukocyte-rich plasma layer and place in a 50 mL tube. Up to 5-10% of the initial RBCs may not have sedimented and thus may still remain in this fraction. This is expected.
5. Wash this fraction once with at least a four-fold volume of appropriate medium. This may require several tubes. Perform a slow spin to remove platelets by centrifuging at 120 x g for 10 minutes at room temperature (15 - 25°C) with no brake. (G to RPM conversion)
6. If excessive platelet contamination is expected, repeat this wash step.
7. Remove supernatant carefully and resuspend the cells in a small volume (typically the nucleated cells from 10mL of blood would be finally resuspended in 0.5 - 1.0 mL of medium).
8. Optional: Any residual RBCs may be lysed with Ammonium Chloride Solution, if desired.

Table 1. Tube Size Recommendations

Volume	Recommended Tube Size
1 - 4 mL	Falcon™ 5 mL polystyrene round-bottom tubes (Becton-Dickinson, Catalog #352058)
5 - 10 mL	Falcon™ 14 mL round-bottom tubes (e.g. Becton-Dickinson, Catalog #352057) OR 15 mL conical tubes (e.g. Corning, Catalog #430053)
> 20 mL	BD Falcon™ 50 mL Conical Tubes (BD, Catalog #352070)

Centrifugation Protocol
Centrifugation may be used to accelerate the sedimentation process.

1. Based on blood sample volume, select an appropriately sized tube (Table 1).
2. Add 1 part HetaSep™ to 5 parts whole blood. Mix well.
3. Centrifuge sample at room temperature (15 - 25°C) at 90 x g with the brake off according to Table 2. (G to RPM conversion)
4. Remove sample from centrifuge and allow to sit undisturbed at room temperature for 10 minutes. This will allow further sedimentation of the RBC and will improve recovery of the nucleated cells.
5. Harvest the leukocyte-rich supernatant into a fresh 50 mL tube. Up to 5-10% of the initial RBCs may not have sedimented and thus may still remain in this fraction. This is expected.
6. Wash this fraction once with at least a four-fold volume of appropriate medium. This may require several tubes. Perform a slow spin to remove platelets by centrifuging at 120 x g for 10 minutes at room temperature (15 - 25°C) with no brake.
7. If excessive platelet contamination is expected, repeat this wash step.
8. Remove supernatant carefully and resuspend the cells in a small volume (typically the nucleated cells from 10 mL of blood would be finally resuspended in 0.5 to 1.0 mL of medium).
9. Optional: Any residual RBCs may be lysed with Ammonium Chloride Solution, if desired.

Table 2. Centrifuge Times Based on Sample Age

		Centrifuge Time (minutes)
Start Volume of Blood* (mL)	Tube Size (mL)	Fresh Blood	24 hr Old Blood	48 hr Old Blood
2	5	1	1	2
3	5	1	1	4
4	5	2	2	5
10	14	5	5	7

*Start volume refers to volume of blood prior to HetaSep™ addition

Note: Contact STEMCELL Technologies' Technical Support at techsupport@stemcell.com for centrifugation speeds and times if processing blood in a 50 mL tube.

References

1. Regidor C, Posada M, Monteagudo D, Garaulet C, Somolinos N, Fores R, Briz M, Fernandez MN: Umbilical cord blood banking for unrelated transplantation: evaluation of cell separation and storage methods. Exp Hematol 27: 380-385, 1999
2. Rubinstein P, Dobrila L, Rosenfield RE, Adamson JW, Migliaccio G, Migliaccio AT, Taylor PE, Stevens CE: Processing and cryopreservation of placental/umbilical cord blood for unrelated bone marrow reconstitution. Proc Natl Acad Sci USA 92: 10119-10122, 1995

A buffy coat suspension is a concentrated leukocyte suspension. It is not mononuclear as the granulocytes are still present. The following brief protocol demonstrates how to prepare a buffy coat suspension from whole blood.

1. Collect whole blood in a blood collection tube containing anticoagulant.
2. Add 1 part PBS + 2% FBS to 1 part fresh whole blood.
3. Centrifuge the sample at room temperature and 200 x g for 10 minutes with the brake off. (G to RPM conversion)
4. Remove the concentrated leukocyte band (this is the buffy coat), and a small portion of the plasma and concentrated RBCs. Platelet contamination in the buffy coat can be decreased by collecting less of the plasma.

Figure 1: Schematic of centrifuged whole blood showing different cell fractions

After centrifugation, one can distinguish a layer of clear fluid (the plasma), a layer of red fluid containing most of the red blood cells, and a thin layer in between, making up less than 1% of the total volume of the blood sample. The thin layer is called the buffy coat (for its buff-colored hue), and it contains most of the white blood cells and platelets.

1. Dilute the whole blood 1:1 with PBS.
2. Layer the diluted sample over a volume of density gradient medium (e.g. Ficoll™, Lymphoprep™) equal to or greater than the initial blood volume.
3. Centrifuge for 30 minutes at 600 x g at room temperature, with the brake off. (G to RPM conversion)
4. Remove and discard the mononuclear cells which form a layer at the Ficoll:Plasma interface.
5. Carefully remove and discard the remaining plasma and density gradient medium.
6. Resuspend the red cell pellet in ACK lysis buffer : completely fill the tube with ACK and incubate on ice for 10 minutes.
7. Centrifuge for 8 minutes at 300 x g.
8. Discard the supernatant and wash the pellet in PBS + 2% FBS.
9. Resuspend the resultant polymorphonuclear cells in appropriate medium (e.g. PBS + 2% FBS).

Ficoll™ is a trademark of GE Healthcare Ltd. Lymphoprep™ is a trademark of Nycomed.

Preparing PBMC from leukapheresis samples by density centrifugation for later cell isolation is time-consuming and requires precision. EasySep™ negative selection kits from STEMCELL Technologies allow you to isolate untouched cells directly from leukapheresis samples, without the need for density centrifugation. This protocol demonstrates how to prepare the leukapheresis samples for downstream cell isolation.

Protocol
If working with large volumes (> 150 mL), first concentrate the leukapheresis cells by centrifuging at 500 x g for 10 minutes. Remove the supernatant and resuspend the cells in 1/10th of the original leukopak volume with recommended medium (e.g. for 300 mL of cells, resuspend in 30 mL of recommended medium).

For small volumes (< 150 mL), add the Ammonium Chloride Solution directly to the cell suspension.

1. Add an equal volume of Ammonium Chloride Solution to the leukapheresis suspension (e.g. for 5 mL of leukapheresis suspension, add 5 mL Ammonium Chloride Solution).
2. Incubate 15 minutes on ice.
3. Centrifuge at 500 x g for 10 minutes at room temperature (15 - 25°C). (G to RPM conversion)
4. Remove the supernatant.
5. Wash the cells by topping up the tube with recommended medium. Centrifuge the cells at 150 x g for 10 minutes at room temperature (15 - 25°C) with the brake off. Carefully remove the supernatant.
6. Repeat the wash step one or more times until most of the platelets have been removed (indicated by a clear supernatant).
7. Resuspend cells at the recommended cell concentration, in recommended medium.

Preparation of a single cell suspension is critical for successful cell isolation. Performing cell separation on samples containing cell clumps will result in cell loss during isolation and may interfere with proper labeling of the target cells.

The following protocols outline how to harvest cells and prepare single cell suspensions from either frozen or primary tissue samples prior to performing cell separation:

Protocol for Frozen Samples

1. Thaw the vial of cells quickly by swirling it in a 37 °C water bath. Transfer thawed cells into a sterile 50 mL conical tube.
2. Rinse the vial with 1 mL of culture medium containing 10 % FBS to recover any remaining cells, and transfer the medium to the new tube.
3. Top up the 50 mL tube with culture medium containing 10 % FBS. Gently invert to mix.
4. Centrifuge the 50 mL tube at 300 x g for 10 minutes at room temperature to collect the cells. (G to RPM conversion)
5. Carefully remove and discard as much of the supernatant as possible, taking care to not disturb the pelleted cells. Gently tap the tube to resuspend the pellet.
6. If cells appear clumpy, calculate the volume of DNase I that should be added to the sample to yield a final concentration of 100 µg/mL. Add DNase I drop-wise to the cell suspension while gently swirling the tube. Incubate for 15 minutes at room temperature.
7. Add 25 mL of culture medium containing 2 % FBS, gently invert to mix and centrifuge as above. Discard as much of the supernatant as possible and gently resuspend the pellet.
8. If cells still appear clumpy, pass the sample through a 30 - 70 µm mesh cell strainer into a fresh conical tube. Rinse the sample tube three times with culture medium containing 2 % FBS and pass through the strainer.
9. The cell suspension is now ready for cell counting and further downstream applications such as cell isolation.

Protocol for Primary Tissue Samples

When working with primary tissue samples, cells must be dissociated from the primary tissue sample in order to prepare a single cell suspension .

1. In a sterile dish containing culture medium and/or dissociation enzyme, mince the harvested tissue. Please contact us or review the current literature for appropriate reagent selection and more detailed procedures for digesting specific tissue types.
2. Pass the sample through a 70 µm or 100 µm mesh cell strainer into a fresh conical tube to further dissociate the cells.
3. Gently pipette 2-3 mL of culture medium over the tissue and cells remaining in the strainer. Repeat this step and then discard the leftover tissue and strainer.
4. Add culture medium - up to 40-50 mL total volume - to the tube and collect cells by centrifugation. Please contact us or review current literature protocols to determine an appropriate culture medium and centrifugation speed for your sample.
5. Carefully remove and discard as much of the supernatant as possible, taking care to not disturb the pellet. Gently resuspend the pellet.
6. (Optional wash step) Carefully add 25 mL of culture medium to the cell pellet, gently invert to mix and centrifuge as above. Discard as much of the supernatant as possible and gently resuspend the pellet.
7. The cell suspension is now ready for cell counting and further downstream applications such as cell isolation.

Cord blood samples can be especially difficult to process as they can have very high platelet counts, leading to clumping even in the presence of anticoagulants.¹ To minimize clumping, samples should be collected in acid citrate dextrose (ACD) or EDTA.¹ If heparin is used as the anticoagulant, 250 U heparin in 1 mL PBS should be added to 10 mL heparin-coated collection tubes. For further tips on how to deal with cell clumps in blood samples, please see Preparing a Single Cell Suspension from Either Frozen or Primary Tissue Samples.

The following protocol demonstrates how to prepare a mononuclear cell fraction from cord blood.

Protocol

Tip: Ensure that the cord blood sample, density gradient medium (e.g. Ficoll™, Lymphoprep™) and buffer (e.g. PBS without Ca⁺⁺ and Mg⁺⁺, and containing 2% FBS) are at room temperature.
1. Transfer the cord blood from the blood bag into sterile tubes or bottle.
2. Remove any visible clumps with a pipette tip. If there are many clumps, pass the sample through a 70 µm cell strainer.
3. Dilute the cord blood with an equal volume of buffer at room temperature and mix gently.
4. Pipette density gradient medium into a sterile tube. For recommended volumes and tube sizes, refer to Table 1.

Table 1. Recommended Volumes and Tube Sizes


Blood (mL)	Buffer (mL)	Density Gradient Medium (mL)	Tube Size (mL)
1	1	1.5	5
2	2	3	14
3	3	3	14
4	4	4	14
5	5	15	50
10	10	15	50
15	15	15	50



5. Carefully layer the diluted cord blood cell suspension over the density gradient medium. It is often useful to hold the tube at a 45° angle during this step, to limit mixing of the density gradient medium and the diluted blood.
6. Centrifuge for 30 minutes at 400 x g at room temperature with the brake off.
7. Remove and discard the upper layer containing plasma and most of the platelets. Harvest the mononuclear cells at the plasma:density gradient medium interface and transfer the cells to a new 50 mL tube.
Tip: Sometimes it is difficult to see the cells at the interface. To ensure good cell recovery, collect some of the density gradient medium along with the cells at the interface. Use a new pipette to remove the mononuclear cell layer to avoid excess platelet carryover.
8. Wash the collected mononuclear cells by adding buffer to about 3X the volume of mononuclear cells or simply by topping up the 50 mL tube. Centrifuge at 300 x g for 10 minutes at room temperature with the brake off.
9. Carefully pipette off and discard the supernatant.
Note: The cell pellet may appear very red due to the presence of red blood cells. The concentration range of immature nucleated red blood cells in cord blood from healthy neonates is 0.03 - 4.8 x 10⁶ cells/mL and, unlike more mature erythrocytes, these cells may remain at the plasma:density gradient medium interface.²
10. Repeat wash step to reduce platelet contamination. Resuspend cells in buffer and centrifuge at 120 x g for 10 minutes at room temperature with the brake off. Carefully pipette off and discard the supernatant. Resuspend cells in desired buffer or medium.
11. Perform a cell count to determine cell concentration.

Ficoll is a trademark of GE Healthcare Ltd. Lymphoprep is a trademark of Nycomed.

References

1. Thomas T et al. "FACS and immunoselection of HSC" in Hematopoietic Stem Cell Protocols (Methods in Molecular Medicine) edited by C.A. Klug and C.T. Jordan. Humana Press, New Jersey. pp 49, 2002
2. Bain B. Normal ranges in neonates, in blood cells: A Practical Guide. Lippincott/Gower, London. pp 155, 1989

Cell counting is an integral part of assessing the results of cell isolation procedures. For best results, both a total nucleated cell count (using 3% acetic acid with methylene blue) and a viability count (using Trypan Blue) should be performed.

Total Nucleated Cell Counts Using 3% Acetic Acid With Methylene Blue

This counting method determines the total nucleated cell count in a sample. The acetic acid lyses the cellular membranes, and the methylene blue stains the exposed nuclei. Because mature red blood cells lack nuclei, they are excluded from this count.

1. Prepare an appropriate dilution of the well-mixed single cell suspension using PBS or serum-free medium. For an accurate representation of concentration, use at least 20 µL of cell suspension to make the dilution.
   
   Example - Preparing a 10-fold dilution
   • Add 180 µL of 3% acetic acid with methylene blue to a microcentrifuge tube or well of a 96-well plate
   • Mix the cell suspension and add 20 µL to the tube or well containing 3% acetic acid with methylene blue
   
   Note: The appropriate dilution will result in a cell concentration that gives 50-100 cells per square in the hemacytometer. We recommend starting with a 10-fold dilution, but the appropriate dilution factor will depend on the approximate number of cells present in the starting sample.
   
   
2. Cover a clean hemacytometer with a clean cover slip.
3. Using a capillary tube or pipettor, apply a small drop (about 10 µL) of the stained, diluted cell suspension to the surface of the hemacytometer at the edge of the coverslip, being careful not to move the coverslip.
4. Place the hemacytometer on the stage of a binocular light microscope, adjust the microscope to 10x power and focus on the cells. Count the cells (stained nuclei) in each of the four outside squares of the hemacytometer (Figure 1), including cells that lie on the bottom and left-hand perimeters, but not those that lie on the top and right-hand perimeters (Figure 2). A minimum of 100 cells should be counted.
   
   
   
   Note: If there are more or fewer than approximately 100 cells per square on the hemacytometer, prepare a new diluted sample using a greater or smaller dilution factor.
   
   
5. Each of the nine major squares of the hemacytometer represents a total volume of 0.1 mm³. Since 1 cm³ is equivalent to 1 mL, the cell concentration can be determined using the following equation:
   
   Total number of nucleated cells/mL = average cell count per square x dilution factor x10⁴
   
   Note: An online hemacytometer calculation tool may be used to easily convert the cell counts into cell concentrations.

Viability Counts Using 0.4% Trypan Blue

Trypan Blue is used to distinguish between live and dead cells. Live cells possess intact cell membranes that exclude dyes such as trypan blue, whereas dead cells absorb/take on these dyes. Therefore the cytoplasm of dead cells stains blue and that of viable cells appears clear. To obtain an accurate representation of cell viability, all reagents, including the cell sample, must be warmed to room temperature before preparing a diluted sample of the cell suspension for counting.

1. Prepare an appropriate dilution of the well-mixed single cell suspension using PBS or serum-free medium. For an accurate representation of concentration, use at least 20 µL of cell suspension to make the dilution.
   
   Example - Preparing a 10-fold dilution
   • Add 80 µL of PBS or serum-free medium to a microcentrifuge tube or well of a 96-well plate
   • Add 20 µL of well-mixed cell suspension to the tube or well containing medium
   • To a fresh tube or well add 20 µL of 0.4% Trypan Blue
   • Add 20 µL of well-mixed diluted cell suspension to tube or well containing 20 µL 0.4% Trypan Blue
   
   Note: The appropriate dilution will result in a cell concentration that gives 50-100 cells per square in the hemacytometer. We recommend starting with a 10-fold dilution, but the appropriate dilution factor will depend on the approximate number of cells present in the starting sample.
   
   Note: To obtain an accurate representation of cell viability, cells should be counted within 15 minutes of staining as cell viability will decrease over time.
   
   
2. Cover a clean hemacytometer with a clean cover slip.
3. Using a capillary tube or pipettor, apply a small drop (about 10 µL) of the stained, diluted cell suspension to the surface of the hemacytometer at the edge of the coverslip, being careful not to move the coverslip.
4. Place the hemacytometer on the stage of a binocular light microscope, adjust the microscope to 10x power and focus on the cells. Count the unstained (viable) and stained (non-viable) cells separately. Count the cells in each of the four outside squares of the hemacytometer (Figure 1 above), including those that lie on the bottom and left-hand perimeters, but not those that lie on the top and right-hand perimeters (Figure 2). A minimum of 100 cells should be counted.
   
   Note: If there are more or fewer than approximately 100 cells per square on the hemacytometer, prepare a new diluted sample using a greater or smaller dilution factor.
   
   
5. To obtain the total number of viable cells/mL in the diluted sample, use the following equation:
   
   Total number of viable cells/mL = (average viable cell count per square) x dilution factor x 10⁴
   
   
6. To obtain the total number of cells/mL in the diluted sample, use the same equation as above, but include both the viable and non-viable cells in the total cell count. Correct for dilution factor as above to obtain the total number of cells/mL in the undiluted aliquot.
7. The percentage of viable cells can be determined using the following equation:
   
   Viable cells (%) =                    number of viable cells                             x 100
                                  number of viable cells + number of non-viable cells
   
   Note: An online hemacytometer calculation tool may be used to easily convert the cell counts into cell concentrations and determine % viability.

The following procedure provides guidelines for isolating cells from samples other than whole blood (e.g. buffy coat sample or leukapheresis) using RosetteSep™, an immunodensity cell separation platform that isolates specific cell subsets during density gradient centrifugation. RosetteSep™ first crosslinks unwanted cells to red blood cells (RBCs) present in the sample to form immunorosettes. When the sample is centrifuged over a density gradient medium (e.g. Ficoll™ or Lymphoprep™), the immunorosettes pellet, leaving highly purified cells at the interface between the plasma and the density gradient medium.

In order to perform RosetteSep™ cell enrichment successfully, it is essential that the sample contain sufficient RBCs; we recommend a ratio of RBCs:WBCs of at least 50:1, and preferably 100:1. This is much more than is commonly found in buffy coat samples.

This is a general procedure; specific conditions may vary according to the cell type being enriched. Find RosetteSep™ protocols for a specific cell type by clicking here.

Sample Preparation
Samples should be resuspended at a white blood cell concentration not exceeding 5 x 10⁷ cells / mL.

Cell Isolation

1. Ensure that the sample, PBS + 2% FBS (phosphate buffered saline + 2% fetal bovine serum), density gradient medium, and centrifuge are all at room temperature.
2. Add RosetteSep™ cocktail to sample and mix well.
3. Incubate 20 minutes at room temperature.
4. Dilute the sample with 2 times the volume of PBS + 2% FBS and mix gently.
5. Layer the diluted sample on top of the density gradient medium using the recommended volumes (Table 1). Be careful to minimize mixing of the density gradient medium and the sample.
   
   

   Table 1. Recommended Volumes and Tube Sizes

   Sample Size (mL)	Tube Size (mL)	PBS + 2% FBS (mL)	Density gradient medium (mL)
   1	5	1	1.5
   2	14	2	3
   3	14	3	3
   4	14	4	4
   5	50	5	15
   10	50	10	15
   15	50	15	15

   
   With 50 mL centrifuge tubes, use a minimum of 15 mL density gradient medium to make it easier to remove the enriched cell layer.

6. Centrifuge for 20 minutes at 1200 x g at room temperature, with the brake off. (G to RPM conversion)
7. Remove the enriched cells from the density gradient medium:plasma interface.
   
   Sometimes it is difficult to see cells at the interface, especially when very rare cells are enriched. To maximize recovery in these instances, remove some of the density gradient medium along with the enriched cells.
8. Wash enriched cells with PBS + 2% FBS. Repeat wash.
9. Use enriched cells as desired. We recommend that residual red blood cells are removed from samples by ammonium chloride lysis to reduce interference with subsequent assays or with flow cytometric analysis. The lysis can be performed as part of either wash step.

Ficoll™ is a trademark of GE Healthcare Ltd. Lymphoprep™ is a trademark of Nycomed.

The following procedure provides guidelines for isolating cells from whole blood using RosetteSep™, an immunodensity cell separation platform that isolates specific cell subsets during density gradient centrifugation. RosetteSep™ first crosslinks unwanted cells to red blood cells (RBCs) present in the sample to form immunorosettes. When the sample is centrifuged over a density gradient medium (e.g. Ficoll™ or Lymphoprep™), the immunorosettes pellet, leaving highly purified cells at the interface between the plasma and the density gradient medium.

This is a general procedure; specific conditions may vary according to the cell type being enriched. Find RosetteSep™ protocols for a specific cell type by clicking here.

1. Ensure that the blood sample, PBS + 2% FBS (phosphate buffered saline + 2% fetal bovine serum), density gradient medium, and centrifuge are all at room temperature.
2. Add RosetteSep™ cocktail to the sample and mix well.
3. Incubate 20 minutes at room temperature.
4. Dilute sample with an equal volume of PBS + 2% FBS and mix gently.
5. Layer the diluted sample on top of the density gradient medium using the recommended volumes (Table 1). Be careful to minimize mixing of the density gradient medium and the sample.
   
   

   Table 1. Recommended Volumes and Tube Sizes

   Sample Size (mL)	Tube Size (mL)	PBS + 2% FBS (mL)	Density gradient medium (mL)
   1	5	1	1.5
   2	14	2	3
   3	14	3	3
   4	14	4	4
   5	50	5	15
   10	50	10	15
   15	50	15	15

   
   With 50 mL centrifuge tubes, use a minimum of 15 mL density gradient medium to make it easier to remove the enriched cell layer.

6. Centrifuge for 20 minutes at 1200 x g at room temperature, with the brake off. (G to RPM conversion)
7. Remove the enriched cells from the density gradient medium: plasma interface.
8. Sometimes it is difficult to see cells at the interface, especially when very rare cells are enriched. To maximize recovery in these instances, remove some of the density gradient medium along with the enriched cells.
9. Wash enriched cells with PBS + 2% FBS. Repeat wash.
10. Use enriched cells as desired. We recommend that residual red blood cells are removed from samples by ammonium chloride lysis to reduce interference with subsequent assays or with flow cytometric analysis. The lysis can be performed as part of either wash step.

Ficoll™ is a trademark of GE Healthcare Ltd. Lymphoprep™ is a trademark of Nycomed.

The following procedure provides guidelines for isolating human cells using EasySep™, a fast, easy and column-free cell isolation system capable of isolating virtually any cell type from any sample source.

This is a general procedure; specific conditions may vary according to the cell type being enriched. Find EasySep™ protocols for a specific cell type by clicking here.

1. Prepare the sample as a cell suspension in the recommended medium, at the concentration indicated on the product information sheet. (Cell Concentration Calculator)
2. Add the EasySep™ cocktail at the concentration indicated on the product information sheet, mix well and incubate at room temperature (15 - 25°C).
3. Vortex the magnetic particles. Ensure that particles are in a uniform suspension with no visible aggregates.
4. Add the magnetic particles to your sample as indicated on the product information sheet. Mix well and incubate at room temperature (15 - 25°C).
5. Bring the cell suspension up to the total volume recommended on the product information sheet by adding medium. Mix the cells in the tube by gently pipetting up and down 2 - 3 times. Place the tube (without cap) into the magnet. Set aside.
6. The magnetically labeled cells will be held inside the tube, while the unlabeled cells can be poured off into a new tube. Separate the fractions as follows (depending on the EasySep™ magnet in use):
• Pick up the EasySep™ magnet, and in one continuous motion invert the magnet and tube, pouring off the desired fraction into a new tube. Hold the magnet and tube in the inverted position for 2 - 3 seconds, and then return to upright position. Do not shake or blot off any drops that may remain hanging from the mouth of the tube.
• Carefully pipette the unlabelled cell fraction into a new tube. Ensure that the pipettor does not touch the sides or the bottom of the original tube.

In negative selection, the desired cells are removed to the new tube. In positive selection, desired cells remain in the original tube. To calculate cell recovery after isolation, use the HIPapp Cell Recovery Calculator.