Functionalized Wire-based Target Cell Isolation: An Ex Vivo Technique to Isolate Cancer Cells from Spiked Peripheral Blood

NOTE: This protocol describes the isolation of HT-29 cells (human colon cancer cell line) from PBS or from artificial mixtures of HT-29 cells and peripheral blood. The same experiment was performed with two additional cell lines (LNCaP and VCaP) and can theoretically be performed with all cells expressing EpCAM.

1. Preparation of Target Cells

1. Cell culture and labeling of cells
   NOTE: In this protocol, cells are cultured in 75 cm² culture flasks. Please adjust the amounts of reagents accordingly if other cell culture devices are used (e.g., 25 cm² culture dishes, 6-well plates, etc.). All liquids used are pre-warmed to 37 °C in a water bath. If not indicated otherwise, all protocol steps are performed at room temperature (RT).
   1. Culture HT-29 cells in McCoy's 5a Modified Medium supplemented with 2 mM L-glutamine, 20 mM Hepes, 10% fetal bovine serum (FBS), and 1% penicillin/streptomycin at 37 °C in a 5% CO₂ atmosphere.
   2. When cells are 90% confluent, remove the medium and rinse the cells with 10 mL of 1x PBS.
   3. To harvest cells, add 2 mL of the cell detachment solution (Table of Materials and Reagents) to the cells and incubate the cells for approximately 5 min at 37 °C.
      NOTE: The detachment solution contains proteolytic and collagenolytic enzymes in 1x PBS, 0.5 mM ethylenediaminetetraacetic acid (EDTA), and 3 mg/L phenol red. Reduce the pre-warming time of the cell detachment solution to a minimum as this will be inactive after 1 h at 37 °C. Cover the whole cell layer to obtain optimal cell detachment.
   4. Check the detachment of cells using an inverted microscope.
   5. Transfer all detached cells to a 50 mL tube prefilled with 10 mL of cell culture medium (same as used in step 1.1.1.) and resuspend the cells by pipetting.
   6. Place the remaining cell suspension on a horizontal roller mixer at RT and proceed to the labeling step.
2. Live cell labeling of target cells
   1. Dilute 1 µL of 5 mM CFSE (supplied in dimethyl sulfoxide, DMSO) in 1 mL of 1x PBS pre-warmed at 37 °C to obtain the 5 µM ready-to-use CFSE labeling solution.
      NOTE: For optimal staining results, use freshly prepared solutions.
   2. Prepare a ready-to-use DNA staining solution (Table of Materials and Reagents) in 1x PBS and store it at 2-6 °C protected from light.
      NOTE: For optimal staining results, use freshly prepared solutions.
   3. Get the cells from the horizontal roller mixer (step 1.1.6.) and pellet the cells at 300 x g for 10 min. Remove the supernatant and resuspend the cells in 10 mL of 1x PBS.
   4. Rinse the cells again with 1x PBS and resuspend the cell pellet in a 500 µL ready-to-use CFSE labeling solution.
   5. Incubate the cells at 37 °C for 15 min and collect the cells after centrifugation at 300 x g for 3 min.
   6. Resuspend the labeled cells in 1 mL of pre-warmed cell culture medium and allow the cells to regenerate at 37 °C for 30 min.
   7. Harvest the cells by centrifugation at 300 x g for 3 min and resuspend the cell pellets in 1 mL of ready-to-use DNA staining solution at 37 °C for 10 min.
   8. Pellet the cells, remove the supernatant, and resuspend the cells in 4 mL of 1x PBS. Assess the cell density using a hemocytometer and check for fluorescence labeling using a fluorescence microscope equipped with 4',6-diamidino-2-phenylindole (DAPI) and fluorescein isothiocyanate (FITC) filters (Figure 1A and 1E).
   9. Centrifuge the cells at 300 x g for 3 min and resuspend the cell pellet according to the cell densities needed for the respective experiment as given in the next section and place on ice.

2. Charging the Wire

NOTE: Cells can be isolated from target cell/peripheral blood spikings at the milliliter scale or by providing a low number of cells at a microliter scale. Whereas the former approach allows for mimicking rare-cell conditions in peripheral blood, the latter may be the preferred way to attach a few cells for the purpose of protocol optimization/testing.

1. Charging the wire using a large volume of cell suspensions
   1. Prepare a 2% BSA blocking solution by dissolving 0.8 g of BSA in 40 mL of 1x PBS (cell culture use). Dissolve BSA by initial vortexing and subsequent incubation at RT on a horizontal roller mixer for 10-20 min.
   2. Rinse empty EDTA tubes/sodium heparin tubes with 2 mL of 2% BSA to remove anticoagulant residues and discard the solution. Block the tube surface by incubation at RT with 5 mL of 2% BSA on a horizontal roller mixer at 15 rpm for 30 min and discard the solution.
   3. Dilute labeled cells (section 1.2.) to a density of 100,000 cells/mL and use 5 mL to charge the wire.
   4. Add 5 mL of the labeled cell suspension (500,000 cells in total) to the tube. Avoid bubbles when adding the cell suspension.
   5. Remove the wire from the storage compartment as well as the rubber cap holding the wire and rinse it in 1x PBS (Figure 2).
   6. Penetrate the rubber tube cap of the EDTA tube (step 2.1.4.) with the help of a syringe needle (20 G) and insert the wire such that the functional part is fully immersed in the cell suspension when the cap is back on the tube and the tube placed on the tilted roller mixer.
      NOTE: The triple-helical functional part of the wire should be covered with cell suspension throughout the charging.
   7. Rotate the tubes on a tilted roller mixer at 5 rpm for 30 min to allow the cells to attach to the wire.
   8. Rinse the wire with 5 mL 1x PBS and store it in the dark at 2-6 °C in a 15 mL tube containing 1x PBS until visual examination.
      NOTE: The functional part of the wire must always be submerged in PBS to avoid harming the cells.
2. Isolating target cells from artificial mixtures with peripheral blood (alternative charging method to: 2.1. Charging the wire using a large volume of cell suspensions) 
   1. Dilute labeled cells (section 1.2.) to obtain cell suspensions with high cell density (> 1,000,000 cells/mL), thereby keeping the volume of cell suspension added to the peripheral blood low.
   2. Add 500 to 500,000 cells to 5 mL of peripheral blood and mix them by inverting the tube.
   3. Remove the wire from the storage compartment, remove the rubber cap holding the wire and rinse it in 1x PBS.
   4. Penetrate a new tube cap with the non-functional part of the device using a syringe needle (20 G) such that the functional part is fully immersed in the spiked blood when the cap is back on the tube and the tube is placed on the tilted roller mixer.
      NOTE: The triple-helical functional part of the wire should be covered with cell suspension throughout the charging.
   5. Incubate the tube on the tilted roller mixer at 5 rpm for 30 min at RT.
   6. Rinse the wire three times in 1x PBS and store it in the dark in a 15 mL tube containing 1x PBS until visual examination.
      NOTE: The functional part of the wire must always be submerged to avoid harming the cells.
3. Charging the wire from low-volume cell suspensions (alternative charging method to: 2.1. Charging the wire using a large volume of cell suspensions)
   1. Resuspend the labelled cells at 1,000,000 cells/mL in 1x PBS.
   2. Fix a 150 mm disposable glass Pasteur pipette horizontally on a rack.
   3. Remove the wire from the storage compartment but keep the yellow rubber cap holding the wire.
   4. Place the wire in the pipette such that the functionalized part is situated in the tip of the Pasteur pipette not touching the glass.
      NOTE: The tip of the wire should not stick out of the Pasteur pipette tip. Avoid plugging the rear end of the Pasteur pipette tip with the rubber cap holding the wire. If attached tight, cell suspensions cannot be loaded from the other side into the pipette tip.
   5. Load 15 µL of the cell suspension into the tip of the Pasteur pipette covering the functionalized part of the wire.
   6. Incubate the wire for 10 min. Manually quarter-turn the assembled wire/Pasteur pipette tip every minute.
   7. Remove the wire from the Pasteur pipette tip, rinse it in 5 mL of 1x PBS and store it in the dark at 2-6 °C in a 15 mL tube containing 1x PBS until visual examination.
      NOTE: The functional part must always be submerged in 1x PBS to avoid harming the cells.

3. Counting Cells on the Wire

NOTE: Always keep the functional part of the wire submerged in 1x PBS to avoid harming the cells.

1. Use a grease pen to draw a rectangle area on a glass slide and add 500 µL of 1x PBS.
2. Bend the non-functional part of the wire and place it on the glass slide such that the functional part is immersed in 1x PBS.
   NOTE: Fit the area of the rectangle and volume of 1x PBS to completely cover the functional part of the wire. Use a double-faced adhesive tape to mount the non-functional part of the wire on the slide.
3. Visually inspect both sides of the wire to enumerate the captured cells (Figure 1B and 1F).
   NOTE: The presence of cells is determined using a fluorescence microscope equipped with filters for DAPI and FITC. Both sides of the wire can be inspected and the number of cells either assessed (for high cell numbers) or counted (only feasible if low numbers of cells are attached to the wire). This step can be skipped if the enumeration of cells is not necessary.
4. After scanning, place the wire back into the 15 mL tube containing the 1x PBS, store the wire in the dark.
   NOTE: Most of the non-functional part of the wire can be cut (including the bend) and removed easing storage.

4. Detachment and Recovery of Target Cells

NOTE: Detachment of the cells shall be done within 4 h after isolation.

1. Dissolve 4 mg of the release buffer component (Table of Materials and Reagents) per mL of 1x PBS and filter the solution through a 0.2 µm sterile filter to obtain the ready-to-use buffer.
   NOTE: The polymer of the wire is composed of a hydrogel which is functionalized with anti-EpCAM antibodies allowing binding to EpCAM-presenting target cells. The release buffer contains a carbon-oxygen proteolytic enzyme capable of cleaving the hydrogel. Proteolytic cleavage results in degradation of the polymer and thus detachment of captured cells.
2. Pre-warm the release buffer at 37 °C for 5 min.
3. Add 1.6 mL of release buffer to fill a 1.5 mL reaction tube.
   NOTE: Tubes designed for 1.5 mL reaction volumes allow application of 1.6 mL which is necessary for submerging the functional part of the wire.
4. Incubate the functional part of the wire in the release buffer at 37 °C (water bath) for 20 min. Use the rubber cap shipped with the wire instead of the tube cap to fix the wire in the 1.5 mL tube to avoid contamination during incubation in the water bath.
5. Transfer the wire/tube assembly to a shaker at RT and 500 rpm for 15 min.
   NOTE: The shaker has an orbit of 1.5 mm.
6. Place the wire/tube assembly in a centrifuge and spin it at 300 x g for 10 min.
7. Remove the wire from the tube, close the cap and centrifuge the tube again at 300 x g for 10 min.
   NOTE: The wire can be stored in 1x PBS at 2-6 °C in the dark for cell counting to assess detachment efficiency/check for cells remaining on the wire.
8. To reduce the volume of the cell suspension, remove all but 100 µL (micromanipulation) or alternatively 300 µL (cytocentrifugation and subsequent laser microdissection) of the supernatant and immediately proceed to single-cell sampling.

Figure 1: Visualizing labeled cells. (A,E) Cells before charging: Cells are labeled with CFSE and counterstained with a DNA intercalating dye showing a range of CFSE (green) signal intensity. (B,F) Labeled cells captured on the wire. (C,G) Wire after cell-detachment procedure. (D,H) Cells detached from wire and recovered by cytocentrifugation onto a slide. CFSE: FITC channel (green). DNA stain: DAPI channel (blue).

Figure 2: Figure 2: Wire and storage compartment. (A) Compartments of the wire. (B) Detail of the functionalized part.