Target Cell Pre-enrichment and Whole Genome Amplification for Single Cell Downstream Characterization

All procedures have been approved by the Ethics Committee of the Medical University of Graz (25-240 ex 12/13). Peripheral blood for spiking experiments was sampled from healthy individuals.

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, experimental data in Representative Results) and can theoretically be performed with all cells expressing EpCAM.

1. Preparation of target cells

1. Cell culture and labelling 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 labelling step.
2. Live cell labelling 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 labelling 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 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 500 µL ready-to-use CFSE labelling 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 labelled 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 labelling using a fluorescence microscope equipped with 4',6-diamidino-2-phenylindole (DAPI) and fluorescein isothiocyanate (FITC) filters (Figure 2A and 2E).
   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 low number of cells at a microliter scale. Whereas the former approach allows for mimicking rare-cell condition in peripheral blood, the latter may be the preferred way to attach 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 labelled 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 labelled 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 3).
   6. Penetrate the rubber tube cap of the EDTA tube (step 2.1.4.) with the help of a syringe needle (20G) 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 labelled 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 (20G) such that the functional part is fully immersed in the spiked blood 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.
   5. Incubate the tube on the tilted roller mixer at 5 rpm for 30 min at RT.
   6. Rinse the wire 3 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 2B and 2F).
   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 enumeration of cells is not necessary.
4. After scanning, place the wire back into the 15 mL tube containing the 1x PBS (see Note of 2.3.6. and section 3.), store it the wire in 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.

5. Single-cell sampling

1. Micromanipulation
   NOTE: Single cells will be added to 2 µL of cell lysis master mix allowing WGA. Prepare master mix according to the number of cells to be processed, calculate extra volumes for loss due to pipetting and place cell lysis master mix on ice.
   1. Prepare the cell lysis master mix (components of WGA kit, Table of Materials and Reagents) according to the protocol outlined by Czyż et al¹⁸. Briefly, mix 2.0 µL of reaction buffer, 1.3 µL of octylphenoxypolyethoxyethanol (10% solution), 1.3 µL of 4-(1,1,3,3-tetramethylbutyl)phenyl-polyethylene glycol (10% solution), 2.6 µL of a proteinase K solution, and 12.8 µL RNase/DNase-free water to obtain a master mix for 10 samples (total volume 20 µL).
      NOTE: For more samples, increase the volumes accordingly. The composition of cell lysis master mix differs from the one used in the alternative procedure utilizing laser microdissection samples (see 5.2.).
   2. Use a grease pen to draw an area holding the cell suspension in place. Adjust area and volume if the cell density is too high (i.e. mark a larger area on the glass slide, load 1x PBS and an aliquot of the cell suspension).
   3. Pipette the entire stained cell suspension (obtained in step 4.8.) onto the glass slide.
   4. Transfer the glass slide to a microscope equipped with a micromanipulator. Let the cells settle for 5 min (Figure 2D and 2H).
   5. Prepare 0.2 mL PCR tubes loaded with 2.0 µL of cell lysis master mix (see step 5.1.1.) and store it on ice.
   6. Collect single cells in 1 µL of 1x PBS using the micromanipulator and transfer the cells into a tube containing cell lysis master mix.
      NOTE: For transferring micromanipulated cells into the cell lysis buffer place the capillary directly into the 2 µL of lysis solution and eject until bubbles can be seen.
   7. Briefly spin down the sample at 2000 x g for 3 s using a desktop microfuge to collect all liquid at the bottom of the tube. Place the samples on ice.
   8. Directly proceed to single-cell WGA (see section 6. and Czyż et al.¹⁹).
2. Laser microdissection (alternative single-cell sampling method to: 5.1. Micromanipulation)
   NOTE: The composition of the master mix used in this section differs from the one used in section 5.1 for micromanipulation. Single cells will be added to 4.5 µL of cell lysis master mix (components of WGA kit, Table of Materials and Reagents) for WGA.
   1. Prepare the cell lysis master mix according to Czyż et al.¹⁹ by mixing 5.0 µL of reaction buffer, 1.3 µL of octylphenoxypolyethoxyethanol (10% solution), 1.3 µL of 4-(1,1,3,3-tetramethylbutyl)phenyl-polyethylene glycol (10% solution), resp., 2.6 µL of a proteinase K solution and 34.8 µL of RNase/DNase-free water to obtain a master mix for 10 samples (total volume 45 µL). Place the master mix on ice.
      NOTE: For more samples, increase the volumes accordingly.
   2. UV-treat membrane-coated slides suited for laser microdissection. Therefore, place the slides in a DNA cross linker device and expose to highest UV for about 10 min.
   3. Assemble the membrane-coated slide in a cytocentrifuge and cytospin the entire cell suspension at 300 x g for 5 min.
      NOTE: When using an in-liquid system, where the cells are cytocentrifuged onto the slide in solution, remove the supernatant after cytocentrifugation and apply a dry-spin at 1140 x g for 1 min.
   4. Directly proceed to laser microdissection or let the cytospins air-dry overnight and freeze the samples at -20 °C for long term storage.
   5. Pipette 4.5 µL of cell lysis master mix into the cap of a 0.2 mL PCR tube and harvest single cells by means of laser microdissection.
   6. Retrieve the PCR tube from the laser microdissection microscope and close the tube. Collect all liquid at the bottom of the tube by short-spin and place the sample on ice.
   7. Proceed with single-cell WGA or store the isolated samples at -80 °C for up to 1 month before further processing.

6. Adapter-linker based whole genome amplification ^18,19

NOTE: Make sure all necessary master mixes (components of WGA kit, Table of Materials and Reagents) are prepared and stored on ice ready to use. Master mixes include the DNA digestion mix 1 for micromanipulated (see 5.1.) samples (containing 2.0 µL of reaction buffer, 2.0 µL of MseI restriction enzyme and 16.0 µL of RNase/DNase-free water) or DNA digestion mix 2 for laser microdissection (see 5.2.) samples (containing 2.5 µL of MseI restriction enzyme and 2.5 µL of RNase/DNase-free water), pre-annealing master mix (containing 5.0 µL of reaction buffer, 5.0 µL of each of the preannealing PCR adapters, and 15.0 µL of RNase/DNase-free water), ligation master mix (containing 30.0 µL of pre-annealed PCR adapters, 10.0 µL adenosine triphosphate solution, and 10.0 µL T4 ligase solution) and primary PCR master mix (containing 30.0 µL of a PCR buffer, 20.0 µL of 10 mM deoxynucleotide mix solution, 10.0 µL of a polymerase mix and 340.0 µL of RNase/DNase-free water). All volumes are given for preparing 10 samples. Adjust accordingly to the number of samples.

1. For cell lysis, place the micromanipulated/laser microdissection samples in a thermal cycler and run single-cell lysis by incubating the cell at 42 °C for 45 min, 65 °C for 30 min and 80 °C for 10 min.
   NOTE: Use a thermal cycler with a heated lid. Cell lysis can be done O/N for 10 to 15 h. In that case, omit the incubation step at 65 °C.
2. Run pre-annealing of the PCR adapters. Therefore, incubate the pre-annealing master mix in a thermal cycler at 65 °C for 1 min followed by further 49 cycles, 1 min each, with progressively decreasing the temperature by 1 °C/cycle. After 50 cycles (at 15 °C), incubate the master mix at 15 °C until further use.
   NOTE: This step is necessary to generate asymmetric adapters suitable for ligation (see step 6.6.) with single-cell DNA subjected to MseI restriction digest.
3. After cell lysis, spin down the lysed samples at 2000 x g for 3 s to collect all liquid at the bottom and place it on ice.
4. To fragment DNA, add 2 µL of the DNA digestion mix 1 to each lysed micromanipulated sample or 0.5 µL of the DNA digestion mix 2 to the laser microdissection sample and run restriction digest. Use a thermal cycler with a heated lid at 37 °C for 5 min followed by a restriction enzyme inactivation step at 65 °C for 5 min.
   NOTE: Digestion at 37 °C may be extended to 3h. This is the only protocol step differing between samples obtained from micromanipulation and samples obtained from laser microdissection.
5. Spin down the samples to collect all liquid at the bottom and place it on ice.
6. For ligating restriction digested DNA and pre-annealed adapters, add 5.0 µL of the ligation master mix to each digested DNA sample and ligate it in a thermal cycler at 15 °C for 1 h.
   NOTE: This step can be extended to 12 h or performed O/N.
7. Spin down the samples to collect all liquid at the bottom and place it on ice.
8. For WGA add 40.0 µL of primary PCR master mix to each of the ligation products and run WGA in a thermal cycler with a heated lid as follows:First, incubate the samples at 68 °C for 3 min. Second, cycle for 15 times at 94 °C for 40 s, 57 °C for 30 s and 68 °C for 1 min 30 s +1 s/cycle. Then add 9 cycles at 94 °C for 40 s, 57 °C + 1 °C/cycle for 30 s, 68 °C for 1 min 45 s +1 s/cycle. Thereafter, cycle for 23 times at 94 °C for 40 s, 65 °C for 30 s, 68 °C for 1 min 53 s +1 s/cycle. Finally, perform a final extension at 68 °C for 3 min 40 s and chill the samples to 4 °C.
9. Spin down the samples to collect all liquid at the bottom and check the DNA quality or store the samples at -20 °C or -80 °C for long-term storage.

7. Quality control of WGA products

NOTE: Check the quality of WGA products based on the range of the DNA smear and the number of amplification products after running a 4plex QC-PCR ⁵. Run WGA aliquots and respective QC-PCR samples on a 1.0% agarose gel for evaluation.

1. Prepare 100 µM stock solutions of all primers used for 4plex quality control PCR (Table 1). Add 8 µL of every primer to 136.0 µL of PCR-grade water to generate 200 µL of primer pool. Vortex the solution for 3 s, spin it down at 2000 x g for 3 s and aliquot 20 µL for storage at -20 °C.
2. Use standard PCR kits to prepare a multiplex PCR master mix containing 6.0 µL of 2x PCR components (except primers), 4.5 µL of PCR-grade water and 0.5 µL of the primer pool.
3. Add 1.0 µL of the WGA products, spin it down and run PCR at 94 °C for 2 min followed by 35 cycles of denaturation at 94 °C for 1 min, primer annealing at 56 °C for 1 min and primer extension at 72 °C for 1 min 30 s and a final extension step at 72 °C for 7 min. Chill it to 4 °C, spin it down and place the samples on ice for gel analysis the same day or at -20 °C for later analysis.
   NOTE: Chilling in the thermal cycler can be performed at 12 °C to increase longevity of Peltier elements.
4. Analyze WGA products (obtained from 6.8.) and respective 4plex QC-PCR products on an agarose gel⁵.