Note: While LUMICKS has experience with these protocols, the outcome cannot be guaranteed and as always, the final decision and responsibility remain with the C-Trap users.

Sample preparation

Cell culture & glass surface functionalization

Cells are cultured using standard protocols and are passaged and plated on glass ~30 mins [1] to a full day [3] prior to the experiment to allow cell attachment. This attachment time varies between cell types and also depends on the glass surface functionalization. Fibronectin [2], or other types of coatings [5] can be used, however, many cell types also naturally adhere well on uncoated glass and may not require a specific surface functionalization [1][3][4]. The level of attachment should be considered carefully as it can affect the cell function and morphology, and is also directly related to membrane tension [2]. In addition, the attachment time can also impact the workflow (see workflow section). The ideal confluency for typical experiments is ~60% which should yield at most a few single isolated cells per bright-field field of view (~110x70 μm).

Plating the cells on 170 μm thick single-use glass coverslips outside of the C-Trap is recommended for cells that require more than ~1h to attach to the glass surface. This subsequently requires a custom chamber assembly (see Fig. 1 and corresponding section). Using 170 μm microscopy-grade glass coverslips will guarantee the best optical performance on the C-Trap. Some polymer coverslip/chambers compatible with 1064 nm light may also be used instead of glass. However, optical performances may not be optimal and there may be a risk of heat-induced damage.

If a shorter than ~1h cell attachment time is enough, the cells can be directly seeded in the standard LUMICKS flow cell (see Fig. 3) despite the flow cell not being gas permeable (fully glass) and the lack of CO2 environment control during the short incubation. The cells can be flowed through the syringe and tubing system of the uFlux and subsequently incubated at 37 °C on the C-Trap with the temperature control [1][6]. This drastically improves experimental control by enabling multiple conditions to coexist on the same flow cell (beads and cells in different channels, buffer exchange, in situ staining, etc.) and makes the workflow generally more reproducible and convenient as it is not impacted by custom chamber-to-chamber variation. The tubing size may also need to be increased to reduce shear stress on the cells flowing in [1], in addition to more extensive cleaning protocols being typically required between experiments.

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Custom chamber assembly and beads

Once the cells are attached to the glass coverslip (and stained or treated, if desired), the custom chamber can be assembled with a standard 1 mm thickness 76x26 mm microscope slide and a ~100 µm spacer (e.g. double-sided tape, see Fig. 1, see example video online. It is critical that the total thickness of the custom chamber remains around 1270 µm for the sample to be compatible with the C-Trap (see C-Trap operations section).Once a custom chamber is built successfully, the solution with beads can be added. Typically, ~2.0 to 5.0 µm polystyrene beads functionalized with concanavalin A are used for this model system [1][4][5].If these are not available, non-functionalized beads can also be used [3] or beads functionalized with other specific proteins [7]. Silica beads can also be used [6] but they will sink faster to the surface and will require the use of slightly more laser power to obtain the same trap stiffness. In terms of size, larger beads are easier for the workflow (easier to trap and move around, easier to interact with the cell)while smaller beads require less laser power to reach the same stiffness, which could reduce laser-induced heating. Typically, a final bead dilution of around 0.001% w/v is used, however, it is recommended to optimize the concentration for your own specific experiment. The ideal concentration should be low enough such that no extra beads interfere with ongoing measurements. The custom chamber’s sides can be sealed with high vacuum grease or other equivalent methods and put into a LUMICKS slide holder with the coverslip on the bottom (see Fig. 1). It is also possible to leave the sides open, but this would accelerate liquid evaporation and likely introduce more sample drift. Beads can also be introduced in situ when using the LUMICKS flow cell or when using a multi-channel custom chamber.

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Cell staining

In some cases, staining for live cell fluorescence imaging is required and can be done just before the experiment. Because the photon detectors of the C-Trap are tuned for single-molecule level signals, the concentration needed to stain the cells is typically lower than recommended by the manufacturer. For example, cells that are stained with ~500 nM SiR-actin for 10 minutes are already sufficiently visible (compared to the recommended 3-4hours at 500 nM). Alternatively, cells can be stained or treated with a different condition in situ if used inside the standard LUMICKS flow cell [6]. It is also possible to achieve in situ change in conditions using custom chambers that have built-in separate channels.

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C-Trap operations

System requirements

A nanosatage is required to perform assays with cells attached to the surface. A m-Trap equipped with a nanostage can also perform these types of assays. However, in the case of the m-Trap, Trap 2 should remain static while the nanostage is used to move the cell towards or away from the Trap 2 bead. In addition, if Trap 1 is not being used, it should be parked far from Trap 2.

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Preparation (custom chamber)

Once the custom chamber assembly is complete, the sample can be used on the C-Trap using a LUMICKS custom slide holder (see Fig. 2 for example holders). It is critical that the total thickness of the custom chamber is as close to 1270 µm as possible (e.g. 170 µm coverslip ~ 100 µm spacer + 1000 µm slide). Indeed, these are the normal dimension used for the alignment of the objective and condenser on the C-Trap and any significant deviation from this can result in incorrect alignment (poor bright-field images and less accurate force detection), and/or collision of the condenser with the custom chamber. In addition, because the condenser requires immersion in oil, the top of the custom chamber must been closed such that an oil droplet can be deposited (i.e. open-top samples are not compatible with the C-Trap system).

1. Turn on the temperature control to 37 °C at least 1h prior to setting up to warm up both the objective and condenser.
2. Lower the objective, add water on the objective, insert the custom slide holder with the custom chamber assembly(Fig. 2). Verify that there is no left-over custom chamber sealing agent (e.g. vacuum grease, or other products, if used) in the working area of the objective. Contact with the sealing agent may damage the objective.
3. Visually inspect that the objective is vertically in line with the region of interest where the cells are plated. If not, adjust the position of the sample using the microstage.
4. Position the nanostage in the middle of its range in x, y, z (100,100,100 µm).
5. While inspecting the z-finder camera, manually raise the objective slightly above the second reflection (i.e. the focus should be inside the chamber but close to the coverslip surface where the cells are plated).
6. While inspecting the moon camera, set the condenser ~50 µm above the usual ‘good’ moon position. This typically means 2-4x more stripes than usual on the moon image (Fig. 2). This will minimize the risk of the condenser colliding with the top of the custom chamber.
7. For this experiment, typically only one trap is needed. After setting up the objective and condenser, change the Trap 1 split to 100%. Be mindful with the overall power used as cells are generally very sensitive. We recommend starting with 5% to 10% overall power(with trap 1 split at 100%), with a resulting stiffness of around ~0.2 pN/nm[1]. If that is not sufficient, it is always possible to increase the overall power, but this may affect the cells. Remember that with Trap 1 split at 100%, there will be no moon image since it is using light from Trap 2.

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Preparation (standard LUMICKS flow cell)

With the standard LUMICKS flow cell, the standard procedure for setting up the objective and condenser can be used. The temperature control should also be turned to 37 °C at least 1h prior to adding the samples into the flow cell. The beads can be loaded into one of the perpendicular channel (e.g. channel 5) while the cells can be flowed through channel 1 (Fig. 3). To reduce the flow shear stress on the cells, the tubing from channel 1 can be exchanged to the larger ~750 µm diameter tubing [1]. Channel 2and channel 3 can be subsequently used for buffer washes, or in situ staining. The cells can be left incubating in the flow cell for several tens of minutes to allow attachment, but for more than 1hincubations, the custom chamber workflow detailed above is recommended instead. Keep in mind that changing the diameter of the tube will affect the flow inside the LUMICKS flow cell. Therefore, any other users using the flow cell for other applications will be affected unless the tubing is swapped back to the original size.

Membrane tethering workflow (steps continue on the next page)

The following steps describe the typical workflow of a cell membrane manipulation experiment while using fluorescence imaging (see Fig. 5 for a visual reference of each step). This workflow is written for experiments with custom chambers but is also valid for the standard LUMICKS flow cell (with slight differences). The membrane tethering workflow is very similar to any type of experiments using a bead to interact with the cell (e.g. experiments on receptor-ligand interactions). For more information about other types of experiments, please see to the last point in the tips & tricks section.

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Step 1: Trap a bead and calibrate. Use the microstage to locate a bead inside the custom chamber. Generally, these can be found floating around in solution or, after some time, sitting on the surface of the channel. When using the standard LUMICKS flow cell, beads can be flowed in and kept separated from the cells (Fig. 3). Once a bead is caught, ensure the bead is located vertically (in Z-direction) about 5x the diameter of the bead away from the surface or more (Fig. 5). For example, if the bead is 3 µm in diameter, calibrate at around 15 µm or more away from the surface. The position of the surface can be approximated by moving the nanostage up in Z until the bead touches the surface (which is visible from a drastic change in force signal and bright-field image of the bead). This surface position may change by several µm’s at different XY positions due to the thermal drift and sample tilt, but it should still provide a good reference. It is useful to save this Z position using a nanostage waypoint. Staying in solution (>5xDiameter)will minimize any near-surface effects that bias the calibration. The bead should also be completely free (not touching any cells, not under any flow),and away from any nearby cell (5-10 µm away in XY) before performing force calibration. Use the hydrodynamic correction (selected by default). It is also recommended to set the forces to zero immediately after calibration. Any subsequent force signals, from various noise/background sources or from real cell interactions can be treated in post-processing.‍

‍Step 2: Bring the cell of interest in plane with the bead. Use the microstage to locate a nearby cell of interest. Consider reducing the microstage speed to avoid losing the bead during stage movement. Locating a cell can be done prior or after step 1 depending on the specific workflow. It is useful to mark the cell’s location with microstage waypoints. Once the cell is located and a bead is brought nearby, the specific contact region of interest of the cell must be moved into the same Z-plane as the trapped bead. This can be done by moving the nanostage up in Z (i.e. moving the cell up), effectively bringing the imaging and trapping planes closer to the cell. Avoid getting too close to the surface as the trapped bead may get stuck on the glass surface

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Step 3: Engage the bead with the cell. Move the bead towards the cell membrane by moving Trap 1 or the nanostage in X and/or Y. By using the nanostage, slight signal changes related to Trap 1’s movement is minimized. It is good practice to start a marker in Bluelake just before moving the bead to capture this initial approach. As the bead is brought closer to the cell, you may notice a background force signal resulting from changes in laser refraction at the sample (due to stage or Trap 1movements, and proximity with other cellular structures). You can verify cell-bead contact by looking at the expected abrupt and large change in force, which should have the opposite sign as compared to the movement (see Fig. 6). E.g. if you are moving the bead slowly to the left (negative movement in X) and begin contacting the cell, the force should push the bead to the right of the trap (large positive force in X). The sign convention on the C-Trap for the Y axis is such that downward movements (and forces) are labelled positive while upward movements (and forces) are negative. In addition, if the bead makes contact, you will likely see the bead change in appearance in the Bright-field camera due to a 3D displacement away from the trap center. It is typically easier to engage the cell when it is spherical compared to when it has a fully stretched and flat morphology. This morphology can be tuned, when desired, through attachment time.

Step 4: Pull a membrane tube. After making contact, you may be able to pull a membrane tube by moving the bead away from the cell. You should also notice a significant difference in force as compared to the initial approach (see. Fig 6, example data). You can verify that it is a real tether by letting the bead go: it should snap back to the cell [1]. It is possible that more than a single membrane tube was pulled. This can be inferred from the force signal if there are visible tether rupture events. Having direct visual confirmation with fluorescence (see step 5 below) is a more reliable and consistent way to check or single-tethers.

Step 5: Optimize the trapping and sample plane for imaging. Some membrane tubes can be observed in the bright-field camera label-free. However, in most cases, fluorescent stains must be used to visualize the membrane with the confocal. Because of the 3D geometry of the cell, the membrane tubes can be pulled from different angles and thus will likely not be exactly parallel to the confocal imaging plane. In addition, the trapping plane may not be aligned with the confocal plane. First, adjust the T1+2z (Fig. 4, step 5) to bring the trapped bead into the focus of the confocal plane. The autofluorescence of the bead (or, if labelled, its fluorescence) will show the largest diameter and sharpest edges when the bead is in focus with the confocal plane. In addition, you should be able to see the part of the membrane tube closest to the bead. This alignment of the trapping plane with the confocal plane using the T1+2z is typically done only once (and may also be done in Step 1). The T1+2z position will then remain aligned with the confocal when using the same bead and same laser powers. Adjusting the T1+2z requires a new force calibration to be performed (as in Step 1) therefore despite having a membrane tube, one should restart the workflow here from Step 1. Once a trapped bead is aligned with the confocal plane, to bring the cell-attached end of the tube into the same confocal plane, the nanostage must be used (depicted as going slightly down in Fig. 4, step 5).

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Tips & tricks

• ‍Ensure that the custom chamber is well-assembled. The spacer should be ~100 µm thick, and it should be well-attached(first) to the slide, and (second) the coverslip. Any loose contact will result in an uneven surface and workflow complications‍
• Use the ctrl+left mouse click shortcut when catching beads to quickly move trap 1‍
• Matching the trapping plane with the confocal plane by moving T1+2z can also be done immediately after catching the first bead, without any membrane tethers being formed, by using the signal from the bead(i.e. it can happen in Step 1). Fine tuning of the different planes can then be done in Step 5‍
• Objects in the confocal focus may not appear in focus in the bright-field image since both modalities have slightly different planes. You may need to adjustT1+2z and the nanostage differently based on whether to prioritize one modality or the other‍
• Releasing the bead from the trap (by shuttering) for a short period of time after cell contact can increase the chances of subsequently forming a tether
• If you have difficulties interacting with the cell despite seeing the overlap between the bead and the cell image, the bead may be too far above the cell and thus not making contact. Move the nanostage up in Z until you observe an abrupt and large change in force (often correlated with a change in the bright-field image of the bead)‍
• If a bead is stuck on the surface or on a membrane, you can increase the overall power to pull it off and decrease it again. However, be mindful that sudden changes in laser power can cause additional thermal drift, and higher overall power may damage the cells. On the next attempt with a new bead, it may also be helpful to reduce the bead’s contact time with the membrane or to change the cell-bead contact position in Z using the nanostage to avoid the bead getting stuck  ‍
• The viability of the cells will decrease overtime on the C-Trap (after ~30 mins to 2 hours), depending on the exact experimental conditions. Be mindful of when that happens in your specific experiments and replace with a new sample when you see the signs. The latter typically include cell blebbing, increased amount of cell debris in solution, and significant changes in cell morphology
• ‍The water on the objective will evaporate faster at 37 °C than at room temperature, typically within 1h to 1.5h‍
• When using the LUMICKS standard flow cell with larger tubing, it is best to keep the pressure to less than 0.10 bar when flushing from the larger tube.‍
• If there is a membrane tether, the bead should be reabsorbed by the cell after the measurement[1]. This is a simple control that can be done after each tether pulling experiment. ‍
• Pulling a tether at different speeds will influence the tether force and which may need to be considered [1]‍
• Other cell-bead interaction experiments can also be performed using a very similar workflow as described here. In the case of a receptor-ligand interaction for example, it is possible to measure the rupture forces at different loading rates and thereby determine the binding kinetics [8]. It is also possible to mechanically trigger the cell and measure its mechanosensing abilities [9]. Several other types of experiments involving beads and cell interactions are possible, from rheology to organelle interactions to motor protein kinetics and much more. See Català-Castro et al. [10] where many of those experiment are compiled in a single review.

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Data analysis

Data format and analysis tools

Several data channels (e.g. Force 1x, Force 1y, Trap 1 position and nanostage position), confocal images and bright-field images are typically useful for cell membrane experiments. From Bluelake, the channels and confocal images are exported to .h5, while the bright-field images are exported to .tiff. Both .h5 and .tiff formats contain all the relevant timestamps information such that they can be correlated in post-processing. Pylake is well-suited for this type of analysis and can be installed following these instructions. The correlation of data channels with bright-field images can be done on Pylake (see ImageStack).The correlation of data channels with confocal images can also be done on Pylake (see multi frame correlation). Alternatively, it is very convenient to use Lakeview to have a quick look at the data channels and the correlated confocal images, to quickly export confocal images into .tiff or to quickly export .h5 data channels into csv format.

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Background force signal  

The trapping laser is refracted when going through cellular structures and this can lead to significant background signals on the force detector when working near or on top of cells. The magnitude of this background signal can be large enough to be mistaken for a real force readout. However, the shape and the sign of the force signal will be very different from a real cell-bead interaction (Fig. 7).Therefore, one must be careful during both the hands-on part to avoid measuring only background signal, and during the data analysis part to avoid mistakenly quantifying the wrong signals.

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References

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[1] De Belly et al. Cell protrusions and contractions generate long range membrane tension propagation, Cell 2023

[2]Pontes et al. Membrane tension controls adhesion positioning at the leading edge of cells, Journal of Cell Biology 2017

[3]Pontes et al. Cell cytoskeleton and tether extraction, Biophysical Journal 2011

[4]Lieber et al. Front-to-Rear Membrane Tension Gradient in Rapidly Moving Cells, Biophysical Journal 2015

[5]Lieber et al. Membrane Tension in Rapidly Moving Cells Is Determined by Cytoskeletal Forces, Current Biology 2013

[6]Evers et al. Single-cell analysis reveals chemokine-mediated differential regulation of monocyte mechanics, iScience2021

[7]Liu et al. Dynamic encounters with red blood cells trigger splenic marginal zone B cell retention and function, Nature Immunology

[8]Yang et al. Molecular interaction and inhibition of SARS-CoV-2 binding to theACE2 receptor, Nature Communications 2020 (Note: AFM reference)

[9]Feng et al. Mechanosensing drives acuity of αβ T-cell recognition, PNAS, 2017

[10]Català-Castro et al. Exploring cell and tissue mechanics with optical tweezers, Journal of Cell Science, 2022

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