Single-cell force spectroscopy

Abstract

The controlled adhesion of cells to each other and to the extracellular matrix is crucial for tissue development and maintenance. Numerous assays have been developed to quantify cell adhesion. Among these, the use of atomic force microscopy (AFM) for single-cell force spectroscopy (SCFS) has recently been established. This assay permits the adhesion of living cells to be studied in near-physiological conditions. This implementation of AFM allows unrivaled spatial and temporal control of cells, as well as highly quantitative force actuation and force measurement that is sufficiently sensitive to characterize the interaction of single molecules. Therefore, not only overall cell adhesion but also the properties of single adhesion-receptor-ligand interactions can be studied. Here we describe current implementations and applications of SCFS, as well as potential pitfalls, and outline how developments will provide insight into the forces, energetics and kinetics of cell-adhesion processes.

Full text

1785
Commentary
Introduction
A broad spectrum of biological processes requires controlled cell
adhesion, including embryonic development, assembly of tissues
and the nervous system, cellular communication, inflammation and
wound healing, tumor metastasis, cell culturing, and viral
and bacterial infection. Although much is known about cell adhesion,
many questions remain unanswered owing to its multiple facets and
complexity. Cell adhesion is commonly defined as the binding of a
cell to a substrate, which can be another cell, a surface or an organic
matrix. The process is regulated by specific cell-adhesion molecules
(CAMs), which are typically transmembrane receptors that comprise
an intracellular domain that interacts with cytoplasmic proteins,
including the cytoskeleton, and an extracellular domain that
specifically binds to adhesion partners (Kemler, 1992). Binding is
commonly heterotypic, but it can be homotypic, such as that
involving cadherins. The major classes of CAMs in mammals include
cadherins, selectins, integrins and Ig-CAMs (cell-adhesion molecules
of the immunoglobulin superfamily). Molecular and genetic
approaches have identified the adhesion proteins and their ligand
specificities, and have determined the processes in which they are
involved. However, the molecular mechanisms by which CAMs
work and how they regulate different types of adhesion are open
debates (Morgan et al., 2007). For example, an extensive array of
proteins is known to be involved in adhesive assemblies, i.e. focal
adhesions [cell–extracellular-matrix (ECM) junctions], but the
contributions of these proteins to the strength of adhesion are not
quantitatively understood (Lo, 2006). To understand cell adhesion,
therefore, the vast amount of qualitative data that is available must
be augmented with quantitative data of the physics of adhesion.
Historically, the strength of the adhesion of a cell to a substrate
has been studied using simple washing assays (Klebe, 1974)
1
.
Surprisingly, given the lack of standardization, washing assays have
proven to be versatile and useful in identifying CAMs, important
ECM components and other proteins that are involved in various
forms of cell adhesion. To estimate the force to which cells are
subjected, various assays that are based on the regulated flow of
media have been implemented, including spinning-disk (Garcia
et al., 1997) and flow-chamber (Kaplanski et al., 1993) assays.
Unfortunately, the shear force that is exerted on the cells in these
assays depends on parameters such as cell size, cell shape and how
the cell is attached to the substrate, and can therefore only be
estimated. For a more controlled and quantitative approach to
measurements of adhesion strength, single-cell methods are needed.
Three types of single-cell force spectroscopy (SCFS) assays have
been developed to measure the strength of cell adhesion down to
single-molecule levels. All three assays use optical microscopes
to observe the cell while force measurements are made, but differ
in how cells are manipulated and forces are determined. The oldest
method uses micropipettes to grasp and hold cells. The detachment
force is measured using a bio-membrane force probe
2
, which can
gauge force between 10
–2
pN (pico-Newtons) and 100 pN (Evans
et al., 1995). A second method uses a pipette to hold a cell while
the strength of interactions between the cell and a functionalized
bead are determined using a laser trap. The laser trap allows three-
dimensional positioning of the bead with nanometer precision and
force measurement from 10
–2
pN to 200 pN (Litvinov et al., 2002).
The third method uses a cell that is attached to a cantilever of an
atomic force microscope (Fig. 1). By combining atomic force
microscopy (AFM) and optical microscopy, cells can be positioned
to assess cellular interactions at a given location on a functionalized
surface, tissue or on another cell (Benoit et al., 2000). The deflection
of the cantilever is used to measure interaction forces. Among SCFS
The controlled adhesion of cells to each other and to the
extracellular matrix is crucial for tissue development and
maintenance. Numerous assays have been developed to quantify
cell adhesion. Among these, the use of atomic force microscopy
(AFM) for single-cell force spectroscopy (SCFS) has recently
been established. This assay permits the adhesion of living
cells to be studied in near-physiological conditions. This
implementation of AFM allows unrivaled spatial and temporal
control of cells, as well as highly quantitative force actuation
and force measurement that is sufficiently sensitive to
characterize the interaction of single molecules. Therefore, not
only overall cell adhesion but also the properties of single
adhesion-receptor–ligand interactions can be studied. Here we
describe current implementations and applications of SCFS, as
well as potential pitfalls, and outline how developments will
provide insight into the forces, energetics and kinetics of cell-
adhesion processes.
Key words: Atomic force microscopy, Cell adhesion, Cellular
interaction, Dynamic force spectroscopy, Extracellular matrix,
Molecular interaction, Single molecule
Summary
Single-cell force spectroscopy
Jonne Helenius
1,
*, Carl-Philipp Heisenberg
2
, Hermann E. Gaub
3
and Daniel J. Muller
1,
*
1
Biotechnology Center, University of Technology Dresden, Germany
2
Max-Planck-Institute of Molecular Cell Biology and Genetics, Dresden, Germany
3
Center for Nanoscience and Applied Physics, Ludwig-Maximilians-University Munich, Germany
*Authors for correspondence (e-mails: jonne.helenius@biotec.tu-dresden.de; mueller@biotec.tu-dresden.de)
Accepted 15 April 2008
Journal of Cell Science 121, 1785-1791 Published by The Company of Biologists 2008
doi:10.1242/jcs.030999
1
In washing assays, poorly or non-adhering tissue-culture cells are washed from a surface
by running a solute (usually medium) over them. The ratio of the number of bound cells
to the number of cells that are initially present provides a measure of adhesion.
2
The bio-membrane force probe is a pressurized red blood cell. The force is measured by
determining the deflection of its membrane.
Journal of Cell Science

1786
approaches, the AFM-based technique allows for the widest
practical force range, from 10 pN to 10
6
pN. This Commentary will
be limited to the AFM-based method, which will henceforth be
referred to as SCFS.
The capability of AFM to image cell topology or characterize
cell-surface properties is outside the scope of this article, and readers
are referred to other reviews (Radmacher, 2002; Dufrene, 2004).
This Commentary will focus on the use of AFM to measure adhesion
strength between a single cell and a substrate that is presented by
a functionalized surface or by another cell. We will explain and
demonstrate the capabilities of AFM and familiarize the reader with
it benefits and limitations.
SCFS set-up and experimentation
Experimental set-up
The basic experimental AFM-type set-up for SCFS is
straightforward. An atomic force microscope that is fitted with a
fluid chamber allows measurements to be made in aqueous
environments under controlled temperatures. Suspended cells are
added to the fluid chamber and allowed to settle. Thereafter, a single
cell is captured by gently pressing a functionalized AFM cantilever
onto it (Fig. 1). This converts the living cell into a probe, which is
brought into contact with functionalized surfaces or other cells at
a set force and for a specific adhesion time. Subsequently, the
cantilever is withdrawn at a constant speed, detaching the cell from
its binding place. During this separation process, the cantilever
deflection, which is proportional to the vertical force that exists
between the cell and substrate, is recorded in a force-distance curve
(Fig. 2). This curve provides the signature of the cell adhesion. The
challenge, however, lies in interpreting this signature, because
various specific as well as unspecific adhesion processes can occur
simultaneously.
Interpreting the cell-adhesion signature
The de-adhesion of a cell from a substrate that is described by the
force-distance retrace curve can be broken into three phases
(Fig. 2). During the initial phase (Fig. 2Ba), the retraction of the
cantilever inverts the force that is acting on the cell from pushing
to pulling. As the overall pulling force increases, the force that is
acting at individual cell-substrate adhesion points increases. If many
receptors act together, the applied detachment force will be
sufficiently high to mechanically deform the cell cortex. The
binding strengths of the receptors, as well as their number and
geometric placement, determines at what force the cell will start
to detach. The largest adhesion force that is recorded, the
detachment force (F
detach
), represents the maximum strength of cell-
substrate binding. Because detachment of the cell is a complicated
process, the maximum adhesion strength represents only a useful
general measure. The work that is required to detach the cell can
also be used to describe the adhesion strength of the cell. It is
calculated from the area that is enclosed by the retraction-
force–distance curve (Fig. 2B). Here, it is important to consider
that the detachment force is a composition of many different
properties of the cell (Bershadsky et al., 2006). These include cell
elasticity, cortex tension, membrane properties, cell geometry and
receptor properties such as binding strength, cooperativity
and placement.
After the cell starts to detach from the substrate, individual force
steps can be observed during the second phase (Fig. 2Bb). During
this phase, the receptor(s) either detaches from the substrate surface
or is pulled away from the cell cortex at the tip of a membrane
tether. While parts of the cell cortex are in contact with the substrate,
either of these processes can occur. During the final phase of
detachment (Fig. 2Bc), the cell body is no longer in contact with
the substrate and, thus, attachment is mediated exclusively by tethers
(Sun et al., 2005a). The force that is required to extend a tether
depends on the lipid composition of the cellular membrane and on
the mechanical properties of the cell cortex. Thus, the lifetime of
a membrane tether is dependent on the receptor-ligand interaction
at its tip, whereas the force that is required to maintain and extend
a tether is not (Marcus et al., 2004). Once initiated, this force is
largely independent of tether length (Hochmuth et al., 1996). In
cell-cell adhesion experiments, retraction distances that approach
100 μm are required, owing to tethers, to fully separate cells (Benoit
and Gaub, 2002; Puech et al., 2006; Thie et al., 1998). There is
ongoing research aimed at trying to use the mathematically tractable
tethers to analyze receptor anchoring (Schmitz et al., 2008). Once
all of the tethers have detached from the substrate, another cycle
of adhesion and detachment is started after a short cell-recovery
time.
A common variation of this method is an inversion of the set-
up. Here, the cantilever of the atomic force microscope is
functionalized with ECM proteins and used to probe immobile
tissue-culture cells (Lehenkari and Horton, 1999). The experimental
set-up is flexible and varies with the biological system. Many
different cells and extracellular adhesion substrates have been used.
It is also possible to apply this approach to detect the molecular
adhesion events of microbial surfaces (Dufrene, 2004). This set-up
can also be used to map the probe-binding properties of cell surfaces.
However, SCFS that uses the cell as a probe has certain advantages
– most importantly, cell-cell interactions can be probed and there
is more freedom in which substrates, such as ECM components,
are presented to the cell.
Versatility of AFM-based SCFS
Many aspects of adhesion can be examined using AFM-based
SCFS, ranging from cell-cell to single-molecule experiments, and
there are no restrictions that govern which CAMs can be studied
or which cells can be used. Initial experiments, which were
performed a decade ago, studied cell-cell adhesion of Escherichia
Journal of Cell Science 121 (11)
A
B
CD
Cantilever
Support
Cell
Fig. 1. Converting a cell into a probe. (A) The apex of a lectin-functionalized
(often by binding concanavalin A) AFM cantilever is positioned above a cell.
(B) The cantilever is gently pushed (generally with a force of <1 nN) for
several seconds onto the cell. (C) The cantilever-bound cell is separated from
the support and allowed to establish firm adhesion. (D) A phase-contrast image
of a cell (arrow) bound to a tip-less cantilever.
Journal of Cell Science

1787
Molecular force spectroscopy of living cells
coli and mammalian cells that were grown on cantilevers, but these
studies were not at the single-cell level (Razatos et al., 1998; Thie
et al., 1998). Shortly thereafter, measurements of the binding
strength between RGD peptides (synthetic peptides that contain
the RGD integrin-binding motif) and osteoclasts indicated the
viability of using living cells and AFM to study single-molecule
binding properties (Lehenkari and Horton, 1999). Since then, a
wide variety of adhesive interactions have been studied using many
types of cells. Table 1 lists the combination of single-molecule
receptor-ligand interactions that have been studied using SCFS.
The various cell types that have been used are also shown in the
table.
Although the atomic force microscope is a high-precision force-
measuring tool, it is versatile. Using piezoelectric actuators, the
atomic force microscope probe can be positioned with sub-
nanometer accuracy at relatively high speeds (>100 μm/second).
With SCFS, cell-substrate contact times can range from milliseconds
to tens of minutes. The imaging capability of the atomic force
microscope can also be used to characterize the adhesion substrate
at a spatial resolution that approaches 2 nm, which clearly exceeds
that of light microscopy (Cisneros et al., 2006; Franz and Muller,
2005; Taubenberger et al., 2007). Thus, for example, AFM images
of cells adhering to aligned collagen matrices revealed cell-induced
rearrangements of individual collagen fibrils (Friedrichs et al.,
2007a). In addition, AFM can be combined with most modern
optical techniques, such as fluorescence-correlation spectroscopy,
wide-field fluorescence, total internal-reflection fluorescence and
confocal microscopy (Chiantia et al., 2007; Franz and Muller, 2005;
Puech et al., 2006; Trache and Meininger, 2005). Commercial
atomic force microscopes that can be integrated into standard and
modern inverted and transmission optical microscopes are
available
3
. In addition, the flexibility and ease of AFM-based SCFS
measurements extends its use from quantitatively characterizing
whole-cell adhesion down to single receptor-ligand interactions.
Examining different aspects of cell adhesion using
SCFS
As one would expect, the strength of adhesion increases with the
length of time a cell is allowed to adhere to a substrate or another
cell. Initially, single receptor-ligand pairs anchor the cell. These
quickly increase in number and undergo modifications to greatly
increase the total strength of adhesion (Friedrichs et al., 2007b;
Taubenberger et al., 2007; Thie et al., 1998). Thus, by simply varying
the cell-substrate contact time during SCFS, both the adhesion
properties of single molecules and whole cells can be quantified.
Observing the adhesion of single molecules
Since its inception (Binnig et al., 1986), AFM has been used to
study molecular interactions (Butt, 1991; Ducker et al., 1991).
However, it took 8 years until the first set-ups that could measure
discrete interactions between single molecules were designed (Lee
et al., 1994; Moy et al., 1994). Since then, this technique, termed
single-molecule force microscopy (SMFS), has been applied to
characterize the binding behavior of many different
oligosaccharides, nucleic acids and proteins (Hansma et al., 2004;
Kedrov et al., 2007; Zhuang and Rief, 2003). Receptor-ligand
interactions are examined by measuring the unbinding (or rupture)
forces between receptors (or ligands) that are attached to the stylus
of the cantilever of the atomic force microscope and ligands (or
I
II
III IV
F
detach
I
II
III
IV
–0.8
–0.6
–0.4
–0.2
0
0.2
0.4
0.6
0.8
86420
Distance (µm)
Force (nN)
Tethers
Jumps
AB
Retract
Approach
(a)
(b)
(c)
PD
Laser
Cantilever
Substrate
3
For example, CellHesion and NanoWizardII BioAFM, JPK Instruments; BioScope II,
Veeco; and MFP-3D, Asylum Research.
Fig. 2. Single-cell force spectroscopy. Depiction of a cell-adhesion measurement (A) for which characteristic approach (green) and retraction (blue) traces are
shown (B). (A) In this technique, the cell and the substrate are brought into contact (AI). The substrate that is probed can be another cell, a functionalized surface or
an organic matrix. The position on a photodiode (PD) of a laser beam (red line) that is reflected off the back of the cantilever measures the deflection of the
cantilever and thus the force that acts on the cantilever. During the approach (denoted by green arrows), the cell (probe) is pressed onto the substrate until a pre-set
force (usually <1 nN) is reached (AII). After a contact time ranging from 0 to 20 minutes, the cell is retracted from the substrate (marked by blue arrows), and a
force-distance curve is recorded (B). This curve corresponds to a cell-adhesion signature. As the strain on the cell increases, bonds that have been formed between
the substrate and the cell break sequentially (AIII) until the cell has completely separated from the surface (AIV). The maximum downward force exerted on the
cantilever of the atomic force microscope is referred to as the detachment force (F
detach
). During the separation of the cell from the surface, two types of molecular
unbinding events can occur. In the first event, the receptor remains anchored in the cell cortex and unbinds as the force increases (denoted as jumps). The second
type of unbinding event occurs when receptor anchoring is lost and membrane tethers are pulled out of the cell. In the unbinding-force–distance curve, long
plateaus of constant force characterize tethers. The shaded area in B represents the measured work of cell detachment from the substrate. The lower-case letters
(a, b and c) denote different phases of cell-substrate detachment (see text for details). Steps I-IV shown in A are also indicated in B.
Journal of Cell Science

1788
receptors) that are immobilized to a surface. Dynamic SMFS probes
these rupture forces at different loading rates (that is, applied force
versus time) to determine the properties of the receptor-ligand
energy landscape (Evans, 1998; Evans and Calderwood, 2007).
These properties typically include the free energy (⌬G) that
separates the bound state from the transition state, the distance
that separates the bound state from the transition state (x
u
) and the
lifetime of the bound state at equilibrium (k
off
). Although these
in vitro measurements provide insights into the behavior of receptor-
ligand interactions, they have limitations.
For example, receptors must be purified, which means removing
them from their biological context, so one cannot be certain of their
functional state. This is of particular concern with integrins that are
known to have several substrate-binding affinity states. Furthermore,
transmembrane receptors are purified in truncated forms, and
therefore consist of only extracellular domains. This is problematic
because it is known that some receptors are regulated through
interactions with cytoplasmic factors, such as the regulation of
integrins by paxillin (Rose et al., 2007).
In contrast to SMFS, SCFS enables single receptor-ligand
interactions to be examined in their cellular environment. Using a
living cell as a probe ensures that the receptors are native. In Table 1,
we list the unbinding forces of receptor-ligand interactions that have
been measured using SCFS. Several of these receptor-ligand pairs
have also been studied using SMFS on solely purified proteins. The
rupture forces that are measured by SCFS and SMFS are generally
in agreement, but some show considerable deviation from each
other. This might indicate that the strength of receptor-ligand
interactions depends on the experimental conditions. Unfortunately,
experimental conditions have not been standardized and are often
not sufficiently documented to allow for rigorous comparisons, i.e.
experiments are performed at different temperatures or the manner
in which the bond is stressed is not clear. Moreover, it is possible
that, although the binding strength that is measured by the two
techniques is similar, the energy barrier that is crossed to break the
bond differs. The external force that is applied to a bond defines
the coordinate along which the bond is forced to break. In SMFS,
receptor-ligand pairs are isolated, truncated and attached to solid
surfaces, whereas in SCFS the receptors are in their cellular
environment. Thus, bonds might break across different paths within
the energy landscape of the binding interaction. Future dynamic
SCFS experiments might show to what extent the biological context
of receptors influences the energy barriers that separate bound and
unbound states.
SCFS is not without limitations – the fact that SCFS uses a cell
as a probe can also complicate certain aspects of single-molecule
measurements. In contrast to the rigid stylus of the atomic force
microscope that is used in SMFS, the applied forces in SCFS cause
the cell to stretch and deform. In addition, because the mechanical
response of cells to deformation is not necessarily linear, dynamic
SCFS measurements are not as straightforward as SMFS
measurements (Evans and Calderwood, 2007). Perhaps more crucial
Journal of Cell Science 121 (11)
Table 1. Receptor-ligand interactions studied by SCFS using living cells as probes
SCFS rupture SMFS rupture
Receptor Ligand(s) force [pN]* force [pN] Cell type Reference
Integrin α2β1 Collagen I and IV 65 (collagen I) Not determined CHO (Taubenberger et al., 2007)
Integrin α4β1 VCAM1 20 Not determined U937 (Alon et al., 2005; Zhang et al., 2004)
Integrin α5β1 Fibronectin 60 (80), 35, 40 Not determined Epithelial, K562 (Li et al., 2003; Sun et al., 2005b; Trache et
al., 2005)
Integrin αLβ2 (LFA-1) ICAM1 35, 40 (80), 70 Not determined Jurkat and (Thie et al., 1998; Zhang et al., 2002;
3A9 HUVEC Zhang et al., 2006)
Integrin αLβ2 (LFA-1) ICAM2 40 (50) Not determined Jurkat (Wojcikiewicz et al., 2006)
Integrin αVβ3 RGD peptide 42 Not determined Bone (Lehenkari and Horton, 1999)
E-cadherin E-cadherin 73 25 CHO (Panorchan et al., 2006b; du Roure et al.,
2006)
N-cadherin N-cadherin 30 Not determined CHO (Panorchan et al., 2006b)
VE-cadherin
†
50 45 HUVEC (Baumgartner et al., 2000; Panorchan et al.,
2006a)
PMN
‡
E-selectin 140 Not determined PMN (Hanley et al., 2004)
PMN
‡
L-selectin 80 Not determined PMN (Hanley et al., 2004)
PSGL-1 (SELPLG) P-selectin 130 150 PMN and LS174T (Fritz et al., 1998; Hanley et al., 2003)
NIH3T3 cell
‡
Concavalin A 80 95 NIH3T3 (Baumgart and Offenhausser, 2003; Chen
and Moy, 2000)
Surface-expressed Concavalin A 86 Not determined NIH3T3 (Chen and Moy, 2000)
mannose residues
Saccharides from blood Helix pomatia lectin 65 Not determined Red blood cells (Grandbois et al., 2000)
types A and O
Galectin 3, galectin 9 Collagen I and laminin 3 Not determined Not determined MDCK (Friedrichs et al., 2007b)
SGLT1 (SLC5A1) Monosaccharides 51 (glucose) Not determined CHO (Puntheeranurak et al., 2006;
Puntheeranurak et al., 2007)
hbhA Heparin 53 50 Mycobacterium (Dupres et al., 2005)
tuberculosis
csA csA 20 Not determined Dictyostelium (Benoit et al., 2000)
discoideum
D-Ala-D-Ala peptide Vancomycin 83 98 Lactococcus lactis (Gilbert et al., 2007)
terminal
‡
The unbinding forces as determined by SMFS are given when known. *The most probable rupture force of the interaction at a loading rate of 1 nN s
–1
is given
when known. Numbers in parentheses are for activated receptors.
†
For these homotypic binding interactions, the adhesion between two cells that express the
same adhesion receptor was probed.
‡
The cell-surface receptor(s) assayed is not known or is not a specific protein. Where two or more rupture forces are given,
different values have been published. PMN, polymorphonuclear cell.
Journal of Cell Science

1789
Molecular force spectroscopy of living cells
is the multitude of possible specific and unspecific cell-surface
interactions. Therefore, special care must be taken to ensure that
the interactions that are recorded occur predominately, if not
exclusively, between the receptor and ligand of interest. To this end,
purified substrates and blocked surfaces are used. Cells can also be
genetically modified to limit the number of possible receptors that
are expressed, and rigorous control experiments that demonstrate
the specificity of the interactions observed must be performed.
The use of a living cell to study single-molecule interactions
at the cell surface has proved fruitful; however, it was shown early
on that SCFS could be used to measure the dynamic-force
spectrum of binding interactions (Chen and Moy, 2000). Making
use of the advantages of living cells, Moy and co-workers
demonstrated that the activation of leukocytes induced changes
in the unbinding-energy landscape of the integrin LFA-1 (ITGAL)
from its ligand ICAM1 (Zhang et al., 2002). Others have since
studied the changes in binding dynamics of various integrins upon
activation by antibodies (Li et al., 2003; Zhang et al., 2004) and
magnesium (Wojcikiewicz et al., 2006). The properties of
genetically modified receptors have also been studied (Alon
et al., 2005).
Studying overall cell adhesion by SCFS
Cellular process, as opposed to single molecules, can be studied
by increasing the cell-substrate contact time
4
. Not surprisingly, in
most SCFS studies the strength of the adhesion between two cells
or a cell and a substrate increases with contact time and contact
area. Generally, retraction curves show that higher detachment
forces are the result of increased numbers of adhesive interactions.
However, significant cell-induced deviations are seen. For
example, the high early adhesion forces that occur between cells
that express the surface receptor Notch and its ligand Delta
diminish as the receptors are cleaved and internalized as part of
the signaling pathway (Ahimou et al., 2004). By contrast, Chinese
hamster ovary (CHO) cells that express integrin α2β1 and are in
contact with aligned type I collagen switch to an activated
adhesion state (Taubenberger et al., 2007). This probably occurs
as a result of the clustering of receptors into load-sharing entities
and not because of the activation of individual integrin molecules.
This phenomenon was observed for integrin-α2β1-mediated
binding of Madin-Darby canine kidney (MDCK) cells to both
type I and type IV collagen (J. Friedrichs, A. Manninen, D.J.M.
and J.H., unpublished). These papers demonstrate that SCFS can
be used to study dynamic and regulated adhesion processes that
occur at the cellular level.
For the assessment of adhesion processes at a level that mimics
the in vivo state, isolated primary cells have been used. For example,
cells that were isolated from zebrafish embryos were used to
examine the importance of non-canonical Wnt signaling for cell
adhesion in early development, and the authors used SCFS to
determine the specific adhesion of different types of primary cells
to functionalized substrates as well as to other cells (Puech et al.,
2005; Ulrich et al., 2005). Moreover, using the same system, the
specific contributions of cell adhesion versus cell-cortex tension to
cell sorting during zebrafish gastrulation have been clarified (Krieg
et al., 2008).
Force-distance curves that trace cell detachment reveal the
unbinding of individual receptor-ligand interactions and, thus,
binding frequency (Gilbert et al., 2007). By simultaneously
measuring the contact area using light microscopy, the number of
active receptors per cell-surface area can be estimated and receptor-
ligand attachment rates found (Gilbert et al., 2007). If combined
with other techniques, such as FACS to determine the number of
particular cell-surface receptors, SCFS allows the fraction
of activated and inactivated receptors to be estimated. A search for
functional states or environmental conditions that tune receptors
might be possible with this combination. Recent SCFS studies on
the adhesion of leukemic cells to bone-marrow stromal cells
showed that the myeloid leukemia fusion protein BCR-ABL
increased integrin β1 expression, which, in turn, increased the
strength of the adhesion between the two cell types (Fierro et al.,
2008). The addition of the anti-cancer drug imatinib mesylate
suppressed the integrin-β1-dependent adhesion to the level of
control cells.
Current limitations and data interpretation
Current SCFS set-ups do have some limitations. Adhesion
measurements that use single cells are time consuming because only
one cell can be characterized at a time. For statistical reasons, many
detachment-force–distance curves must be recorded, which limits
the length of the contact times that can reasonably be assayed.
Furthermore, the almost unavoidable thermal drift in AFM
complicates long-contact-time experiments (>20 minutes) and the
tight adhesion of cells after longer contact times (>1 hour) exceeds
the capability of the system
5
. Thus, SCFS is currently restricted to
short contact times that range from milliseconds to ~20 minutes.
There is also a high cost associated with SCFS; fortunately,
inexpensive atomic force microscopes that have been specifically
developed for SCFS should soon become available.
As with most new techniques, SCFS needs to mature. Presently,
enthusiasm to publish comes, at times, second to rigorous
examination of the data. Of particular concern are: (1) the need to
establish controls that demonstrate the specificity of the molecular
interaction being studied; (2) the temptation to over-interpret
numerical data that are gleaned from unverified mathematical
models; and (3) the difficulty in appreciating the complexity of
both the physics and biology of the systems studied. This situation
will improve as familiarity with the technique increases and
standard experimental procedures and data-analysis norms are
adopted.
Perspectives
The use of SCFS is still in its infancy and has much potential for
development. This potential is based on the versatility of SCFS
and the enormous variety of cell biological and medical applications
to which it can be applied. Here, we have demonstrated that SCFS
provides a ‘force signature’ of the cell-adhesion process and have
shown how this tool has been used to study CAMs and the
dynamics of regulated adhesion processes in living cells. In
practice, all possible forms of cell adhesion can be studied, with
limitations that include the restriction of experiments to short
contact times and the high associated cost, as discussed above. The
combination of AFM with advanced light-microscopy imaging has
yet to be applied to its full advantage, and studies in which force
4
To study single-molecule unbinding, contact times from 0 to 0.5 seconds are generally
used. The rate at which bonds form depends on the system that is assayed.
5
At high force, the adhesion between the cantilever and the cell is weaker than that between
the substrate and the cell. Using concanavalin A to immobilize the cell to the cantilever,
forces of up to 50 nN can be measured before cells detach from the cantilever. This force
is, however, dependent on cell type.
Journal of Cell Science

1790
measurements are correlated with changes in cell shape and
structure will emerge in the foreseeable future. The development
of single-cell force spectroscopes that have standardized cell-
handling and analysis routines will also provide a possibility to
expand the experimental parameters that can be addressed with
this innovative technique.
We thank A. Taubenberger, C. Franz, J. Friedrichs, M. Krieg and
K. Simons for stimulating discussions. The Deutsche
Forschungsgemeinschaft, the European Union, the Munich Center for
Integrated Protein Science and the Volkswagenstiftung supported this
work.
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