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Tumor Processing Protocol

 

 

 

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mdacc

 

 

WebMaster ~ Tammy Davis
Last Updated ~ March 30, 2011

 
Tumor Processing & Marker Staining for Marker Analysis Protocol

 Reagents

  • Cell culture medium*
  • Phosphate Buffered Saline (PBS)
  • Accumax/EDTA (Innovative Cell Technologies, AM105)
  • Histopaque (Sigma, 10771)
  • Staining Buffer (see No. 14)
  • Lineage-depletion kit (Miltenyi, 130-090-858)
  • Antibody of interest conjugated with a dye
  • Appropriate antibody control
  • Cell culture medium without serum
  • Heat-inactivated, undiluted serum

* Most PCa xenografts are cultured in IMDM+15% FBS. Du145 is cultured in RPMI+7% FBS

Procedure

  1. Weigh the tumor in a pre-tared sterile 40 mm petri dish.
  2. Add a small volume (1-2 mls) of cold culture medium to the petri dish and chop it into small pieces using a razor blade (Surgical Blades, No. 23). Further mincing can be done using dissecting scissors to obtain a thick soup of tumor cells/pieces.
  3. Transfer the soup to a 50 ml tube using a cut 1-ml pipette tip. Add ~20x the volume of cold medium, invert to mix and spin at 900 rpm for 5 minutes. The tissue pieces and dissociated cells should pellet down while most of debris should remain in the supernatant
  4. Suck off the supernatant and add 30 mls of cold PBS. Invert to mix and spin again at 900 rpm for 5 minutes. Discard supernatant.
  5. Resuspend the cells/chunks in Accumax/EDTA solution at a concentration of 10 mls/0.5g of tissue. Incubate at room temperature (RT) for 30 minutes on a shaker. The digestion reaction should never be done at 37°C as the high temperature inactivates the enzymes. Accumax should be stored cold at all times (unopened stock at -20°C).
  6. At the end of the incubation, briefly vortex the tissue solution and then set the tube straight for about 2 minutes to allow the largest chunks to settle to the bottom. Transfer the supernatant containing the dissociated cells to a fresh tube and spin at 900 rpm for 5 minutes.
  7. While the cells are spinning, prepare a 40µm nylon mesh by placing it on a 50 ml tube and gently pre-wetting it with medium. At the end of the spin, discard the supernatant and resuspend cells in a small volume of medium (1-3 mls). Slowly, pass the cells through the mesh without applying too much pressure. If the mesh becomes clogged, replace with a new, pre-wetted one and continue until all the cells have gone through. Only single cells or doublets should pass through the mesh (30 µm nylon mesh can be used if too many doublets or aggregates exist). Wash the mesh 3x with medium.
  8. Count the total cell number. Live cells can be distinguished from dead cells using a viability dye such as erythrosin B (ATCC, 30-2404) at a 1:1 ratio of cells to dye. If the cells are very concentrated, a 1:10 dilution can be done to achieve a more accurate count. The following is an example of a calculation of cell number.

      Total volume of cell suspension = 10 mls

10µl of cell suspension added to 90µl of media (total dilution = 10x)          10µl of diluted solution mixed with 10µl of erythrosin B (total dilution = 20x)          Load 10µl of this mix onto hemocytometer and count the cells in the 4 squares at 4 corners

               Cell number = no. in squares x 10,000 x 20 (total dilution) x 10 (total volume)   ( 4 squares)

            The viability % is live cells  x 100 (live + dead cells)

  1. The live cells are separated from dead cells, debris, and red blood cells through use of a histopaque-1077 gradient. Briefly, the cell suspension volume is calibrated at ~1-1.5 million total cells (dead + live) per ml. In a 15 ml tube, add 3 mls of histopaque. Carefully load the same volume of cell suspension upon the histopaque so that 2 layers are clearly observed. The loading should be done with a 1 ml pipette tip as a larger pipette tends to disrupt the interface between the two layers. Multiple tubes can be used depending on the total cell number. Alternately, a 50 ml tube can also be used with 15 mls of Histopaque followed by 15 mls of cell suspension at a concentration of no more than 1 million/ml (dead+live). If an extremely large number of cells is obtained through digestion (e.g. >50 million live cells), then only a portion of that number should be used for further processing as the flow cytometry time and speed limit us to analyzing a maximum of ~20 million cells.
  2.  Spin the cells on Histopaque at 400g for 30 minutes at RT.
  3.  At the end of the spin, an opaque layer of cells should be observed at the interphase of the two layers. These are the live cells. The dead cells and debris should pass through the Histopaque layer and collect at the bottom of the tube. Using a 1 ml pipette tip, carefully remove the live cell band and transfer to a new tube. Combine the live cells from all tubes and spin at 900 rpm for 5 minutes.
  4.  Resuspend in 1-5 mls of medium and count the cell number the same way as  above. The cell number obtained after Histopaque should be no less than 75% of the original number.
  5.  Spin again at 900 rpm for 5 minutes.
  6.  To deplete the xenograft tumor cell population of mouse (host) lineage-positive cells, a lineage-depletion kit (Miltenyi, 130-090-858) is used. The kit is a magnetic labeling system for the depletion of mature hematopoietic cells such as T cells, B cells, monocytes/macrophages, granulocytes and erythrocytes, as well as some mouse stromal (fibroblasts, smooth muscle, and endothelial) cells. For this purpose, the cells are magnetically labeled with a cocktail of biotinylated antibodies against a panel of so-called ‘lineage’ antigens (CD5, CD45R, CD11b, Gr-1, and Ter-119), and anti-biotin beads. Mouse cells can also be removed using anti-H2Kd antibodies (Pharmingen) that recognize the mouse histocompatibility class I antigens. If primary human tumors are used, a human-specific lineage-depletion mix (either custom-made through some companies or directly obtaining from commercial sources) should be used. This depletion mix has antibodies to several markers associated with normal human leukocytes, endothelial cells, mesothelial cells, and fibroblasts (CD2, CD3, CD10, CD16, CD18, CD31, CD64, and CD140b). In either case, the procedure is done at 4°C in the dark. All solutions should be pre-cooled. The staining buffer is prepared as follows:  sterile PBS supplemented with 0.5% bovine serum albumin (Sigma, A-4161; can also be replaced with FBS) and insulin (Sigma, I-6634; 100x stock is 0.5 mg/ml, final is 5 mg/ml). The insulin stock, once prepared, is good for one month at 4°C. The volumes given below are for up to 5 million cells. When working with higher numbers, scale up the reagent volumes and total volumes.
    • Suspend cell pellet in 40 ml of buffer
    • Add 10 ml of biotin-antibody cocktail
    • Mix well and incubate for 10 minutes at 4-8°C in dark
    • Add 30 ml of buffer
    • Add 20 ml of anti-biotin microbeads
    • Mix well and incubate for additional 15 minutes at 4-8°C in dark
    • Wash cells by adding 10-20x labeling volume and centrifuge at 900 rpm for 5 minutes. Pipette off supernatant completely
    • Resuspend up to 107 cells in 500 ml of buffer
    • For the separation, assemble the MACS apparatus by attaching the magnet on the stand and placing the column (Miltenyi, 130-042-201) in the magnetic field. Prepare the column by rinsing with 500 ml of buffer. Make sure that there is a 15 ml collection tube below the column.
    • Apply cell suspension to the column. Allow the cells to pass by gravity and collect the effluent as fraction with unlabeled cells, representing the enriched lineage-negative cell fraction.
    • Wash the column with 500 ml buffer 3x.
    • Spin down the effluent at 900 rpm for 5 minutes and discard supernatant
    • The pellet contains the lineage-negative tumor cell population. At this juncture, the cells should be resuspended in medium and another count should be done to obtain the final cell number.
  1. Spin one more time at 900 rpm to bring the cells in 110 ml of cold staining buffer (volumes given are for 5 million cells; scale up for higher numbers). Remove 10 ml from cell suspension and add to 90 ml of buffer in another tube. This is to set aside some cells for the control staining.
  2. Add 10 ml of the antibody of interest (such as CD44-FITC) to the 100 ml cell suspension, mix by tapping, and incubate at 4°C in the dark for 15 minutes. The tube should be gently tapped every 5 minutes to prevent the cells from settling down. For the control tube, use 1 ml of appropriate isotype control (for example, CD44-FITC is a mouse IgG2b , k. Therefore, the control would be FITC-mouse IgG2b , k isotype control immunoglobulin). If no control Ab is available, then no antibody should be used as the negative control.
  3.  Add 10 mls of cold staining buffer and spin at 900 rpm. Resuspend in cold serum-free medium (+1% antibiotics) at a concentration of 2 million cells/ml. To hand over to the FACS facility, the cell suspension should be transferred to 5 ml polystyrene tubes at no more than 2.5 mls/tube. Collection tubes should also be prepared using 5 ml polypropylene tubes containing 1 ml of FBS (+1% antibiotics). This provides a cushion for the cells when they come through the flow machine at high speeds. Tubes should be labeled “+” or “–“ depending on cell number expected. For example, for CD44 sorts, since the number of “+” cell are much lower than the “–” °cells, we generally prepare 1 “+” tube and 3 “-” tubes.
  4. Add 7-AAD (Molecular Probes, A-1310) to the sort tube 10 minutes before analysis. This is a viability dye that only penetrates cells with compromised cell membranes, much like erythrosin B. The stock concentration of 7-AAD is 100 mg/ml (100x, final = 1 mg/ml).
  5. All reagents and samples must be turned in for FACS analysis on ice. FACS is carried out anywhere from 1000-2000 cells/second, depending on the size and aggregation properties of the cell type used.
  6. Once FACS analysis has been carried out, the “+and “–” cells should be spun at 1000 rpm, resuspended in 0.5 – 3 mls of regular culture medium, and counted. The sorted cell number reported by the FACS machine is usually inaccurate and it is important to make an actual count as most of the experiments depend on a direct comparison between equal numbers of  “+” and “–” cells.
  7. At the end of every sort, it is generally a good idea to do a post-sort analysis as a quality control. This is best done by re-running the sorted cells through the flow cytometer and determining the purity. However, if the number of sorted cells is too low, then to conserve the sample, it would be better to simply look at a drop of the suspension under the inverted fluorescence microscope and determine the percent purity. Some dyes (eg. PE), however, are not easily visible under the microscope and a post-sort can only be done using flow. It should also be noted that during the sort, it is possible that many antigens get knocked off cells because of high speed collisions making the purity percentage for the “+” population appear much lower than it actually is. This can be overcome by plating the cells on coverslips for 2-4 hours, thereby allowing them to recover some of their antigen expression before re-staining them to analyze for purity.
  8. In vivo injections are done either subcutaneously or orthotopically into the prostate. For subcutaneous (SC) injections, cells should be suspended in serum-containing medium at the appropriate concentrations and mixed with an equal volume of matrigel (1:1 ratio). Note that matrigel is in liquid form only at 4°C and should be kept on ice at all times. For SC, the volume for each injection should not exceed 100 ml (including matrigel). For surgical orthotopic implantation(SOI), the cells are also resuspended in regular medium. However, the volume should not exceed 20 ml per injection as the mouse prostate is tiny and injection of higher volumes would cause it to rupture. If the cells are being injected with matrigel, the matrigel concentration should not exceed 25% of the total volume (5 ml matrigel + 15 ml cell suspension). If fibroblast support cells are being injected with the epithelial cells, they should be mixed at a ratio of 2.5:1 fibroblasts to epithelial cells beforehand. For example, if 100K cells are to be injected with fibroblasts and matrigel, the injection would consists of 15 ml of cells/fibroblasts (100K cells + 250K fibroblasts) + 5 ml matrigel.