An Overview of Single-cell Force Spectroscopy
This note gives a broad overview of single-cell force spectroscopy from the main concepts to the existing measurement methods and the main applications.
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What is Force Spectroscopy?
Introduction to Force Spectroscopy
The presence of the adhesion phenomenon in many fields like cell biology, mechanobiology, biophysics, and biomedical engineering, has led to the development of tools to quantify adhesion forces from the nano- to the micro-scale such as single-cell force spectroscopy.
Force spectroscopy describes methods that measure interactions and forces between molecules, cells or particles. To quantify such force, most methods measure the behavior of a molecule when it is exposed to stretching or torsional (twisting) forces. More specifically, the molecule will have one end bound to a surface and the other to a force sensor, whose displacement is measured to determine the forces.
A range of tools have been developed to accurately manipulate molecules and measure subsequent interactions, including atomic force microscopy (AFM), optical tweezers, magnetic tweezers, and micropipettes.  These tools have contributed greatly to our understanding of interactions that span many orders of magnitude, from protein-DNA interactions at sub-nanometer scale to cell-cell and cell-substrate interactions at the micrometer scale.
The video below illustrates simply the working principle of force spectroscopy to measure the interaction forces between a probe - here the FluidFM probe and cells adhering to a surface.
A full webinar dedicated to single-cell force spectroscopy realized in collaboration with Nanosurf is also available here. In the next section, various methods that can be employed to perform AFM-based single-cell force spectroscopy, will be introduced.
Existing methods to perform single-cell force spectroscopy
At the cellular scale, understanding and quantifying cell adhesion is important for our understanding of a wide spectrum of biological processes, such as differentiation, proliferation, and migration. Additionally, AFM gained popularity for SCFS because it brought about the first tool with high measuring resolution and a broad range of detectable forces. Prior tools, like micropipette-aspiration and optical tweezers, were highly precise but were limited by either low force resolution or low detectable forces. This has led to the development of assays for single-cell force spectroscopy (SCFS), wherein binding forces between two cells or between a cell and a molecule of interest are measured.
The introduction of AFM-based techniques for SCFS opened doors for studying adhesion in live cells in near-physiological conditions. AFM-based techniques comprise the non-exhaustive list of the following techniques:
Single-cell force spectroscopy with standard cantilever
In the direct force spectroscopy approach, the cell to be analyzed, can be either adhering to a surface and subsequently approached by a standard cantilever to study the interactions; or, oppositely, the cell can be attached onto the cantilever and employed as a probe to approach the surface, or a cell adhered onto the surface, to investigate cell-cell interaction in native environment.
In the first case, despite a rather straightforward experimental approach, limitations exist due to the shape and materials properties of commercially available standard cantilevers. Indeed, most of standard AFM cantilevers present a sharp tip which can damage the cell deposited on the surface upon approaching and probing. In addition, force measurement performed in liquid environment with standard cantilevers can be quite challenging due to a number of potential disturbances due to air bubbles, contaminants or electrostatic forces interfering with the cantilever approach or retraction.
In the second possibility, traditional AFM-based SCFS relies on the attachment of the living cell to the AFM cantilever via chemical functionalization.  This method presents a number of drawbacks. Firstly, adhesion measurements with AFM-based SCFS are time consuming and low-throughput. To achieve firm adhesion of the cell to the cantilever for force measurements, the cantilever must be functionalized first. This chemical functionalization of the cantilever “glues” the cell of interest to the cantilever irreversibly , which means that each cell requires a separate functionalized and calibrated cantilever, a process that can take 30 min . This approach is thus low throughput, with a complex and time-consuming cell attachment/detachment methodology. Importantly, the process of immobilization could also alter cell physiology by affecting the cell surface.
Secondly, the force range of measurements with current AFM-based SCFS is limited by the force with which the cell is bound to the cantilever, meaning that that only forces below the coupling strength of the cell to the force sensor can be measured. This approach is thus intrinsically limited to measurements in the early phases of cell adhesion formation, unable to provide the kinetics of adhesion forces at longer timescales (i.e. mature cell-cell contacts) [5, 6]. Importantly, the irreversible immobilization of a cell also limits biological replicate measurements in SCFS because of the long time required to perform a statistically significant number of measurements. This laborious method requires specific expertise and may also damage the cell or denature the cell surface. 
The difficulties related to the attachment of a cell onto a cantilever and the limitations of adhesion force resolution obtained with standard cantilevers led scientists to investigate alternative methods such as the colloidal probe technique to perform single-cell force spectroscopy.
Single-cell force spectroscopy with colloidal probe technique
The colloidal probe technique was developed to investigate interaction forces between single colloids and a substrate . Instead of a cantilever with a sharp AFM tip, a colloidal particle is glued to a tipless cantilever, and the subsequent cantilever is approached to the cell seating on a surface. This technique allows the researcher to gain a higher force sensitivity due to the broader surface approaching and probing the substrate in solution. The rounded shape and the material of the particle attached to the tipless cantilever ensures a smoother approach to the cell. However, this technique is also affected by several inherent limitations caused by the need to glue the colloid to the AFM cantilever and by the fact that the particle being broader than a standard cantilever, more settling time and care are required upon the approach.
A partial solution was obtained with the use of the FluidFM technology to attach reversibly the colloid particle via negative pressure, removing the problems encountered with the gluing step. In 2013, Dorig et al. (2013) used FluidFM technology to study single-colloid interaction with a substrate in liquid environment, demonstrating the use of FluidFM as a reliable and versatile alternative to colloidal probe AFM.  The authors use FluidFM to reversible attach colloids via negative pressure, measure adhesion to the substrate of colloids, followed by their release via a short surge of positive pressure. The exchange of colloids, thanks to the reversible nature of their attachment to the FluidFM probe, allows for quick variation of different colloids within the same experiment and the investigation of long-term interactions of colloids on surfaces. FluidFM enables analyzing mechanical properties like adhesion in a time- and cost- efficient manner, thereby accelerating research and new product development.
In the next part, another stage was reached with the FluidFM technology, for the reversible and smooth immobilization of a single cell, to study cell-cell or cell/substrate interactions.
Single-cell force spectroscopy enabled with the FluidFM technology
Unlike previous techniques, with the FluidFM OMNIUM, live cells can be reversibly immobilized onto the cantilever by applying and maintaining adequate negative pressure of the fluidic channel on the FluidFM probe. This gentle physical, rather than invasive chemical, immobilization allows SCFS measurements to be done in native conditions (without chemical treatment of cells). With this ability, FluidFM probes can be used multiple times, hence improving the experimental reproducibility. In addition, once the measurement of a single cell is completed, the FluidFM probe can be controlled to absorb another cell for further measurements.
The FluidFM technology thus increases both throughput and efficiency of SFCS measurements, drastically reducing the time required to obtain statistically relevant data compared to conventional AFM-based SCFS. 
Features and benefits of single-cell force spectroscopy measurement with FluidFM
Direct and semi-automated force measurement
Through a simple and reversible cell immobilization
Broad force range
Measure forces from pN up to µN!
Measure up to 200 cells a day! Isolate single-cell, accurately and reliably.
Applications of single-cell force spectroscopy
In the following, single-cell force spectroscopy demonstrates its potential in a broad of biological applications and studies.
Single-cell force spectroscopy in eukaryotic cells
Physical studies of single cells allow insights into biophysical and mechanobiological phenomena in differentiation, growth, and proliferation. In cancer research, immunology and neuroscience, the mechanical properties of cells and their interactions with their environment such as with other cells (cell heterogeneity) or the properties of biological structures and surfaces are key parameters. Likewise, for implant materials there is a clinical need to understand and control how various cells adhere to it.
Many studies have now used FluidFM technology to investigate adhesion behaviour in a broad range of model cell types, including HeLa cells, HEK cells, HUVEC cells, C2C12 cells, and oocyte cells [7-10] and to investigate other aspects of cell adhesion, such as adhesion forces in mature cell-cell contacts , or the effect of electric current on cell adhesion.  We highlight a few studies here.
Potthoff et al. (2012) were the first to establish serial quantification of adhesion forces of different cell types using FluidFM-based SCFS.  Figure 1 shows the experimental principle of SCFS using FluidFM technology. Potthoff and colleagues were the first to show the pace of SCFS accelerated to up to 200 yeast and 20 mammalian cells per probe when replacing the conventional cell trapping cantilever chemistry of AFM with FluidFM technology. This way, statistically relevant data could be recorded in a rapid manner and a range of different cell types could be analyzed with one single probe. The authors performed adhesion measurements for yeast Candida albicans and for the most widely used mammalian model cells, HeLa cells, and for HEK cells. Monitoring adhesion of mammalian cells revealed mean adhesion forces of 600 nN of HeLa cells on fibronectin and were one order of magnitude higher than those observed for HEK cells. The use of FluidFM technology also allowed authors to perform long-term dynamic measurements of adhesion of cells to substrates for the first time, as the traditional AFM-based SCFS is not amenable to long-term measurements due to the need for chemical immobilization of cells for measurements. This property means that FluidFM technology allows not only for quantitative and statistically relevant data in a rapid manner but also makes it possible to monitor the temporal changes in long-term adhesion processes that are relevant for biofilm or tissue formation.
In 2017, Sankaran et al., used FluidFM technology to investigate the adhesion interactions between single cells (mouse myoblast C2C12 cells) and different types of substrates: two types of RGD-presenting surfaces (one non-covalent and one covalent surface).  They used C2C12 single living cells attached to the FluidFM probe to perform approach-retract movements on the substrates and record force curves. The results obtained from force curves provided quantitative insight into adhesive interactions between cells and substrates by providing information on the unbinding between the cell and the surface such as adhesion force, detachment distance, and binding energy.
Single-cell force spectroscopy in microbials cells
Understanding microbial adhesion can have important implications for infection biology and for biomedical applications, such as in providing insight into the role of biofilms in the pathogenesis of implant infections. The term biofilm refers to groups of microbial cells that adhere to the exopolysaccharide matrix present on the surface of medical devices.  Microbes first attach onto biomaterials, after which intercellular interactions between proteins lead microbes to cluster together, forming microcolonies and biofilms. Microbial adhesion is thus essential to biofilm formation, survival, and pathogenesis of microbes. By measuring adhesion, biofilm formation can be studied and addressed in the further optimization of surface materials, like in the design of implants as well as anti-microbial or non-fouling surfaces.
Studies looking to investigate the mechanisms that underlie adhesion interactions in single microbial cells have also benefited from FluidFM-based SCFS. Single microbes can be easily immobilized to the FluidFM probe by applying negative pressure and adhesive interactions between the microbes and substrates or other microbes can be measured. Mechanical studies with FluidFM therefore give valuable insights for microbial research, extending the body of knowledge gained through classical biological research methods. In 2015, Potthoff and co-workers showed that the FluidFM probe can be used for reversible immobilization of single bacterium under physiological conditions then used for subsequent measurements of adhesion forces.  The use of FluidFM allowed the authors to study bacterial adhesion for contact times of more than 2 hours, whereas previous AFM-based SCFS measurements at the time had recorded interaction forces after a maximum of 60s.  The reversible immobilization enabled by the FluidFM also allowed, for the first time, to use a single probe for serial measurements on different bacterial cells, enabling direct comparison of cell-to-cell variation. This allowed authors to quantify contact time and setpoint dependence of the adhesion forces for morphologically different bacteria species, the Gram-negative rod-shaped model microorganism E. coli and the clinically relevant spherical Streptococcus pyogenes. Because the latter grows in chains, FluidFM was also used to study the sequential detachment of bacteria out of a chain, revealing distinct force patterns in the detachment curves of the bacteria. This was the first study to demonstrate the potential of the FluidFM technology for quantitative bacterial adhesion measurements of cell-substrate and cell-cell interactions relevant in biofilms and infection biology. Importantly, the authors also show that bacteria still divided after SCFS measurements using the FluidFM, highlighting that bacteria survive the measurements and thus that the FluidFM can thus be used for multiple experiments.
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