The purpose of the feasibility of biopsy viability is to ensure that cells remain alive following biopsy. Not all experiments will require cells to stay alive following biopsy. For example, experiments that require only one biopsy per cell. In these cases, the feasibility of biopsy viability can be skipped.

Viability becomes critical when biopsies are collected from the same cells at different timepoints, as biopsies can rarely be collected from dead cells. It therefore becomes important to preserve cell viability as much as possible, which this step of the experiment aims to do. Carrying out the feasibility of biopsy viability also gives an indication of how many cells statistically need to be targeted at T0 in order to be able to rebiopsy X number of cells at T1. Like the feasibility of biopsy collection, the feasibility of biopsy viability only needs to be performed once for each cell line, meaning that once viability rate after biopsy in the cell line and the best parameters are known, the feasibility can be skipped and the clean biopsy collection can be carried out directly.

Prerequisites

This workflow is intended to be carried out immediately following the define piercing parameters section of the feasibility of biopsy collection (see Define Piercing Parameters in Experimental Procedure in Feasibility of Biopsy Collection), where 20 cells have been biopsied and their positions recorded.

Materials

• Cell media
• Sytox Green (or equivalent non-endpoint viability die) 

Define Piercing Parameters

Defining the piercing parameters should be carried out as described in (see section Define Piercing Parameters in Experimental Procedure in Feasibility of Biopsy Collection) but with an additional action as Step 2. of the Rinsing:

1. Select a control cell and run observation:
2. 1. Save the location of a control cell
   2. 1. Return to the location of the cell just biopsied.
      2. Select a cell close by that looks like one you would biopsy.
      3. Mark the cell by opening the Observation workflow, select the cell and save it to a group ('ControlXX', where XX is the equivalent number of the biopsied cell). 
   3. Run an observation: 
   4. 1. Select Position, by selecting all groups to select all points that have been created so far.
      2. Observe imaging parameters: (as defined for brightfield in Imaging parameters in Cell Line Onboarding)
      3. 1. Action: Record image
         2. Cantilever position: Retract
         3. Preset: imaging preset (as defined in Imaging parameters in Cell Line Onboarding)
         4. Iterations: 1
         5. Time interval: Doesn't matter
         6. Focus: Restore focus

For more information on the Observe workflow see Observe - FluidFM OMNIUM (online manual).

Observation

The observation workflow in ARYA allows you to set up an automated workflow to take images of your saved points (biopsy locations) at specified intervals using specified imaging settings. For more information on the Observe workflow see Observe- FluidFM OMNIUM Manual (online manual). To run the observation workflow:

1. Following all extractions, run the water wash as described in Wash Plate Preparation (see Experimental Preparation in Feasibility of Biopsy Collection) and dry the probe (see Probe Drying in Buffer Entry, Aspiration and Deposition Test section in Experimental Procedure for the Feasibility of Biopsy Collection). Do not remove any of the plates or the probe from the instrument.
2. Add 0.5 mL of sterile ddH2O, to compensate for evaporation during extraction every 3 hours that the lid is open
3. Place the cover on the dish (to prevent evaporation during observation) carefully without moving it (to prevent offset in the saved points in ARYA).
4. Switch on the heating (to 37°C) and CO2 control (to 5%) in the OMNIUM.
5. Set both plates in the system to observation mode.
6. Set up an observation workflow:
7. 1. Select Position:
   2. 1. Select all groups to select all points that have been created
   3. Observe:
   4. 1. Action: Record image
      2. Cantilever position: Observation height (or as determined in Imaging parameters in Cell Line Onboarding)
      3. Preset: Brightfield observation settings (as determined in Imaging parameters in Cell Line Onboarding)
      4. Iterations: XX ( = (Desired Time (min)/Time Interval (min))+1)
      5. Time interval: MM mins (as determined in the section Seeding Density and Tracking Timing-Interval in Cell Line Onboarding)
      6. Focus: Restore focus
      7. Choose a well and retractfor parking (Recommended: drying station)
      8. Click 'Observe'

Viability Assessment

1. When the observation iterations are finished, add 1 mL of media to the well, containing 2 µL of Sytox Green.
2. Incubate for 10 minutes.
3. Set up an observation workflow:
4. 1. Select Position: 
   2. 1. Select all groups to select all points that have been created
   3. Observe:
   4. 1. Action: Record image
      2. Cantilever position: Observation height (or as as determined in Imaging parameters in Cell Line Onboarding) 
      3. Preset: Brightfield observation settings and fluorescent green observation settings ( as determined in Imaging parameters in Cell Line Onboarding) 
      4. Iterations: 1
      5. Time interval: Not important, can be set to any number
      6. Focus: Restore focus
      7. Disable the 'merge imaging' option
      8. Choose a well and retract for parking (Recommended: drying station)
      9. Click 'Observe'

Cell Tracking Tool

The tracking tool allows you to track cells across multiple images. It only tracks cells in images gathered in the Observation workflow in ARYA and the tool was developed to aid manual tracking (doesn’t yet incorporate image recognition). Here, you can click on the cell in each image to track its movements over time. You can also tag events in the cell’s life at certain images in the sequence e.g., splitting, detaching, dying etc. When you save the tracked cell’s path, you can then return the probe to the last marked position of the cell. A video of the tool can be seen in Figure 1. 

Figure 1. Video showing how a cell is tracked using the tracking tool in ARYA. 

To track the cells following the extended observation and viability test:

1. Open the tracking tool by clicking on the icon:
2. Select the group and the point used during the extraction
3. Localise the biopsied cell in each image:
4. 1. If the cell is alive and visible, mark it by clicking on it
   2. If the cell has split, detached or died, use the annotation tool to record this:
   3. 1. If the annotation does not exist yet, 'add an entry'
      2. If the annotation has been previously created, select from the dropdown menu.
5. Click 'save changes'.
6. The last image (fluorescent) should show whether the cell remains alive or has died. Record the condition of the cell. Cells displaying the following criteria should be considered ‘dead’:
7. • Cell exhibiting green fluorescence (Sytox Green positive)
   • Cell is no longer present or cannot be located
   • Cell has detached from the substrate
8. Repeat Steps 2. - 5. for all biopsied cells and all controls.
9. Compare the viability of the control cells and the biopsied cells.

Volume Tool

The volume measurement tool allows the users to quantify the amount of extracted cytoplasm that is deposited in a droplet and ultimately processed for downstream analysis (e.g., RNA sequencing). It is also useful at the feasibility stage to ensure that the volumes that you are able to extract from a particular cell type will be enough for downstream analysis.

• During the extraction, one image is acquired when the probe is inserted into the cell, and another image after the extraction is finished and the cantilever retracts to exit the cell (see Step 3 of Extraction in Define Parameters in Experimental Procedure for the Feasibility of Biopsy Collection, and Figure 2 below).
• The user is requested to focus on the cantilever so that the buffer/oil or biopsy/oil interface is visible.
• By comparing the images before and after the extraction, the user can calculate the extracted volume with the volume tool.

The individual extractions need to be stored in a point group so that the data can be retrieved after the extraction experiment is completed.

Figure 2: Example images before and after a single cell biopsy

To use the volume calculation tool, open the volume measurement tool in ARYA by clicking on the icon:

Several areas will appear (numbered areas correspond to the labels in Figure 3):

1. Point Group Selection – Here, you select the group your cell of interest belongs to. Once the group is selected, the individual points (extracted cells coordinates) will appear. By selecting one point, the corresponding data acquired during the extraction will be displayed.
2. Image Panel – Images acquired during one extraction experiment are displayed twice (on the left and right).
3. Volume Guide - Appears superimposed on the cantilever.
4. Volume Selection Icons – The volume can be selected in 3 different ways (explained further below).
5. Properties – On the right-hand side, the user can adjust properties of the cantilever.
   Note: Always use the real channel height when calculating the volume for accurate calculation: the default value is set to 1 µm, whilst the nominal value of a nanosyringe is 1.5 µm. For the measured value (wafer specific), contact the Support indicating the Wafer LOT number.

Figure 3. Different areas of the volume estimation tool. 

To calculate the volume:

1. Select an image on the left-hand side that corresponds to the focused cantilever image before the extraction.
2. Select on the right-hand side the image that corresponds to the focused cantilever image after the extraction.
3. A differential image will appear where the user can visualize the interface before and after the extraction (see Figure 4). The volume within the two interfaces is the extracted volume.
   
   Figure 4. Differential image of the cantilever before extraction (orange arrow) and the cantilever after extraction (red arrow). Note the volume in the center of the cantilever between the two interfaces that corresponds to the extracted volume. 
4. Adjust the volume guide over the cantilever by selecting the first icon in the upper right corner (see Figure 5). The volume guide is an indication of volumes that can be aligned on top of the cantilever (see the blue guide marked with volumes over the cantilever in Figure 4). By clicking and holding the volume guide the user can move it around and align it to the cantilever.
   
   
   Figure 5. Volume selection icons. The first icon on the left allows you to adjust the position of the volume guide on the cantilever.
5. The other three icons in Figure 5 are three different methods of outlining the volume. Select the one that you believe is the most appropriate for your calculation:
6. 1. Fast calculation without the preload – By selecting the second icon, a red line will appear that can be moved up and down, sliding it over the volume guide (see Figure 6). By adjusting the red line to the line of the interface, the user can get a quick estimation of the volume. Keep in mind that this is not the exact volume but a close estimate.
      
      Figure 6. Fast calculation without the preload.
   2. Fast calculation of the extracted volume – Similarly to the icon, a red rectangle will appear that can slide along the volume guide. The upper line can be adjusted to the height of the preload interface and the lower to the height of the extracted interface (see Figure 7). This gives an estimation and not the exact volume.
      
      Figure 7. Fast calculation of the extracted volume.
   3. Lasso icon – By selecting the lasso, the user can trace a more precise area between the preload interface and extracted interface, compared to the previous options (see Figure 8). With the mouse left-click hold the lasso and trace the area. With the mouse right-click hold and trace the areas within the previously selected area, to exclude possible pockets (areas without the extract). This gives the most accurate volume measurement. Pillars account for roughly 15 femtoliters of volume (estimated over 8 pillars for channel height of 1.5 µm) and can be either removed or left in.
      
      Figure 8. Lasso icon. 
      Note: all calculations can also be done on single images (bright field or fluorescent acquisitions), without the differential image. 

To store the data and export it:

1. “Set Volume” on the lower right corner will link the calculated volume to the point by displaying it next to the point name.
2. “Save Result” on the lower left corner will display a tick to confirm the image used to calculate the volume is stored in the point group, with the red overlay of the selected volume (see Figure 9). The femtoliter volume will also be saved in the image metadata.  
   
   Figure 9. Setting the volume and saving the result.  
3. The process can then be repeated for each point or cell.
4. After all cells from a given group have been analysed, exit the tool by pressing “Cancel” (set volumes that are not saved prior to this step, won’t be stored in the point group).
5. The volume data can then be exported from the side bar, the gallery, or the point group. By selecting “Export” a dialog will appear where the user can select both image and data options.

Note: selecting multiple result images from the gallery view will generate a single .csv file with one selected point per line, with the corresponding calculated volumes in femtoliters.

Criteria for Continuing to Clean Collection

The purpose of the feasibility studies is to ensure that the chosen cell type can be biopsied in the FluidFM OMNIUM as required for the planned clean experiment. To this end, we recommend that the experiment can continue when:

• Experimental set-up parameters are known and confirmed (e.g., cell seeding density, imaging parameters, time intervals for imaging etc.)
• Buffer is known or confirmed experimentally to be compatible with the biopsy workflow
• The piercing parameters for the cell line have been defined, and cells can be biopsied with minimal issues under all experimental conditions.
• A reasonable amount of extract can be taken from the cell (we normally recommend at least 0.5 pL)
• Biopsied cells do not have a significantly different viability from non-biopsied cells in the FluidFM OMNIUM (only necessary if cells are to be biopsied more than once, this threshold can also be lowered at the testers discretion, however it may require a higher number of extracts at T0 in order to have the desired number of biopsies at T1).