Characterization of the specific interaction between the DNA aptamer sgc8c and protein tyrosine kinase-7 receptors at the surface of T-cells by biosensing AFM
Characterization of the specific interaction between the DNA aptamer sgc8c and protein tyrosine kinase-7 receptors at the surface of T-cells by biosensing AFM
Michael Leitner 1 2
Alexandra Poturnayova 0 1
Constanze Lamprecht 1 2
Sabine Weich 1 2
Maja Snejdarkova 0 1
Ivana Karpisova 1
Tibor Hianik 1
Andreas Ebner 1 2
0 Institute of Biochemistry and Animal Genetics, Slovak Academy of Sciences , Moyzesova 61, 900 28 Ivanka pri Dunaji , Slovakia
1 Faculty of Mathematics , Physics, and Informatics , Comenius University , Mlynska dolina F1, 842 48 Bratislava , Slovakia
2 Institute of Biophysics, Johannes Kepler University Linz , Gruberstrasse 40, 4020 Linz , Austria
We studied the interaction of the specific DNA aptamer sgc8c immobilized at the AFM tip with its corresponding receptor, the protein tyrosine kinase-7 (PTK7) embedded in the membrane of acute lymphob l a s t i c l e u k e m i a ( A L L ) c e l l s ( J u r k a t T- c e l l s ) . Performing single molecule force spectroscopy (SMFS) experiments, we showed that the aptamer sgc8c bound with high probability (38.3 ± 7.48%) and high specificity to PTK7, as demonstrated by receptor blocking experiments and through comparison with the binding behavior of a nonspecific aptamer. The determined kinetic off-rate (koff = 5.16 s−1) indicates low dissociation of the sgc8c-PTK7 complex. In addition to the pulling force experiments, simultaneous topography and recognition imaging (TREC) experiments using AFM tips functionalized with sgc8c aptamers were realized on the outer regions surface of surface-immobilized Jurkat cells for the first time. This allowed determination of the distribution of PTK7 without any labeling and at near physiological conditions. As a result, we could show a homogeneous distribution of PTK7 molecules on the outer regions of ALL cells with a surface density of 325 ± 12 PTK7 receptors (or small receptor clusters) per μm2.
DNA aptamer; PTK7; T-cell; Single molecule force spectroscopy; Energy landscape; Molecular recognition; Recognition imaging
Cancer is a major societal challenge and its detection
and identification at the earliest stage are crucial for
efficient and successful treatment. Acute lymphoblastic
leukemia (ALL) is a common type of blood cancer. It is
characterized by aggressive and uncontrolled division of
abnormal lymphocytes, which spread to various parts of
the body and penetrate and destroy healthy body tissue
. Rapid identification and classification of the
pathogenic cells is important for choosing the correct therapy
for ALL patients. Conventional diagnosis comprises a
combination of methods, including morphologic,
cytochemical, cytogenetic, or immunologic tests [2, 3], as
well as bone marrow biopsy . Additional techniques
to further classify the type of leukemia include flow
cytometric immunophenotyping  and polymerase
chain reaction studies [4, 6, 7]. A novel approach that
may render these elaborate and invasive procedures
unnecessary is based on the recognition of cancer-specific
biomarkers on the surface of cancer cells by DNA/RNA
Aptamers are synthetic short single-stranded DNA or
RNA oligonucleotides that fold into unique
threedimensional shapes. These structures enable highly
selective and specific targeting of molecules with high
affinities comparable to those of antibodies. The small
size and rather simple structure of aptamers relative to
antibodies makes them easier to be synthesized and
chemically modified. Moreover, they display low to no
immunogenicity among other advantages. Therefore,
aptamers have emerged as a new molecular tool in
clinical medicine to detect and isolate proteins, and to act
as targeting and therapeutic agents [9–12]. The DNA
aptamer sequence sgc8c has been synthesized to
specifically recognize ALL T-cells , where it is known to
bind with high affinity (Kd = 0.8 ± 0.09 nM) to the
protein tyrosine kinase-7 (PTK7) [14, 15]. PTK7 has also
been found to be overexpressed in various other cancer
types, including colorectal cancer and cancers of the
lung, prostate, lymph nodes, and breast [16–19]. Thus,
sgc8c has become a promising conjugate for targeted
delivery of chemotherapeutics [20–22], photothermal
agents [23, 24], immunotherapeutics , and contrast
agents [26–28], and for noninvasive diagnosis  of
Recently, O’Donoghue et al. addressed the first step of
sgc8c mediated cancer cell targeting on a single
aptamerreceptor level using atomic force microscopy (AFM) .
The aptamer was linked to the tip of the AFM cantilever
and brought into contact with the plasma membrane of
HeLa cells. In their proof of principle experiment rupture
forces of 46 ± 26 pN between sgc8c and PTK7 on the cell
surface were measured only at one given force load and
showed that the binding strength of aptamer and antibody
to cancer cells was about equal under these setting. Here,
we expand on this work, and include dynamic aspects of
the molecular recognition between sgc8c and PTK7 on
Jurkat T-cells by conducting single molecule force
spectroscopy (SMFS) under variation of the force load. We
performed AFM recognition imaging to gain data on the
distribution of PTK7 receptors on Jurkat cells. SMFS has
become an increasingly popular technique in the
development of new pharmaceuticals to explore the interaction of
new therapeutic molecules with cell membranes and whole
cells [31–33]. The technique enables determination of
energetic, thermodynamic, and kinetic parameters that
describe the free-energy landscape of the interacting ligand
target molecule complex [34, 35]. In particular, SMFS
yields the dissociation rate constant (koff) and the width
of the interaction potential (xβ), which characterize the
microscopic basis of bond formation of a ligand–receptor
pair . In this study we employed SMFS to measure
the relative off-rate, the main determinant of the affinity
between sgc8c aptamer and PTK7 receptor complexes. In
addition, biosensing AFM also enables localization of
binding sites and their distribution on cellular surfaces
by using ligand-functionalized tips in analogy to those
used in SMFS for high-resolution AFM imaging [31, 37,
38]. Here, we employ the well-established method of
simultaneous topographic and recognition imaging
(TREC), which has been successfully applied to localize
binding sites on isolated molecules [39–42], artificial
, and native membranes , as well as whole cells
[45, 46] in the past decade. We performed AFM
recognition imaging to probe the distribution of PTK7
receptors for the first time on the single receptor level on
Materials and methods
All chemicals were used in their highest available
purity. PBS and HBSS buffer salts, acetic acid, and citric
acid were obtained from Sigma-Aldrich (Vienna,
Austria), cell media and HEPES buffer were purchased
from PAA (Pasching, Austria), and Cell-Tak from BD
Biosciences (Erembodegem, Belgium). In all
experiments only ultrapure MilliQ (MQ) water (Millipore,
Darmstadt, Germany) with 18 MΩ resistance was used.
T-cell culture and preparation
TIB 152 cells (Jurkat clone E6-1, ATCC, Wesel,
Germany) were cultured in RPMI 1640 medium
containing 10% fetal calf serum (FBS) supplemented with
1% penicillin/streptomycin and 1% HEPES buffer and
maintained in an incubator under air atmosphere with
5% CO2 at 37 °C. Cells were passaged twice a week
and reseeded at a concentration of 1:5. For AFM
experiments, cells were used 3–4 d after splitting and
immobilized on round glass slides (diameter 22 mm,
VWR, Vienna, Austria). For cell attachment, glass slides
were first cleaned with 80% isopropanol and ultrapure
sterile MQ water (Millipore, Darmstadt, Germany) and
dried. Next, the surface was coated with BD Cell-Tak in
5% acetic acid (Sigma-Aldrich) by hand-spreading the
solution with a 50 μL glass micropipette and left for
air drying under the laminar flow. After rinsing with
70% ethanol and a final washing step with sterile MQ
water, cells were added. For this the cells were
centrifuged and the cell pellet resuspended in 1500 μL RPMI
media without FBS, supplements, and phenol red. Five
hundred μL of cell suspension was transferred to each
Cell-Tak coated glass slide and incubated under air
atmosphere with 5% CO2 at 37 °C for 30 min. Cell
density, their condition, and adherence were checked under
the light microscope. Before chemical fixation, cells
were rinsed three times with PBS buffer (137 mM
NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM
NaH2PO4, pH 7.4) to remove media components, then
treated with 4% formaldehyde in HBSS (Vienna, Sigma
Aldrich) over a period of 60 min at room temperature,
washed again in PBS three times, and used immediately
or stored in the fridge for a maximum of 5 d.
Preparation of sgc8c aptamer AFM sensors
The sequence of the PTK7 specific DNA aptamer sgc8c was:
5′ (SH- or) NH2- ATC TAA CTG CTG CGC CGC GAA AAT
ACT GTA CGG TTA GA-3′. For the specificity proof the
nonspecific DNA aptamers TDO5 was used: 5′ NH2- CAC
CGG GAG GAT AGT TCG GTG GCT GTT CAG GGT CTC
CTC CCG GTG-3′. The sequences of the aptamers were taken
from the paper by Huang et al. . Both aptamers were
purchased from Thermo Fischer Scientific GmbH
(Darmstadt, Germany). All aptamer solutions were prepared
by dissolving lyophilized oligonucleotides in TE buffer
(1 mM EDTA, 10 mM Tris, pH 8).
TREC measurements were performed with magnetically
coated cantilevers MACLever Type VII (Keysight, Santa
Rosa, USA). SMFS experiments were done with MSCT
probes (Bruker, Karlsruhe, Germany). For both TREC and
SMFS experiments the identical cantilever functionalization
protocols were applied in which DNA aptamers were tethered
to the apex of silicon(nitride) tips using a distensible
heterobifunctional poly(ethylene) glycol linker. As silicon
(nitride) is known to be an inert material, the tip was first
chemically activated by deposition of APTES from the gas
phase to ensure a sufficient number of reactive sites, but
without forming a 3D network of the functionalization agent that
would cause variations in the unbinding length of rupture
events. For this, silicon (MACLevers) or silicon-nitride
(MSCT) tips, respectively, were amino-functionalized
according to the gas-phase deposition protocol published previously
. In brief, chloroform-cleaned cantilevers were placed in
an argon filled 5 L desiccator together with a vial filled with
60 μL freshly distilled amino-propyl-triethoxysilane (APTES,
Sigma-Aldrich, Vienna, Austria) and another vial with 20 mL
trimethylamine (TEA, Sigma-Aldrich, Vienna, Austria), and
allowed to react for 120 min. Then the desiccator was flushed
with argon gas for 5 min and left for 48 h for curing process.
For tethering the aptamers, either the NHS-PEG-acetal linker
 was employed to covalently couple amine terminated
aptamer NH2-sgc8c to the AFM tip or the NHS-PEG-PDP
linker  to bind thiol terminated aptamer SH-sgc8c.
Attachment of the reference aptamer TDO5 was done
In the case of NH2-terminated aptamers, freshly APTES
coated tips were incubated in a chamber containing a solution
of 1 mg NHS-PEG-acetal linker dissolved in 500 μL
chloroform. Thirty μL TEA was added as catalyst. After 120 min
reaction time, tips were washed with chloroform and ethanol
(three times for 5 min.) and dried again in a gentle nitrogen gas
stream. To obtain the aldehyde function, tips were immersed
for 10 min in 1% citric acid solution, washed three times in
water, and dried under N2 gas. The aldehyde functionalized
tips were immersed in ~40 μL PBS solution containing 1 μM
NH2-sgc8c aptamer and 2 μL freshly prepared aqueous
solution of 1 M NaCNBH3 (Sigma-Aldrich, Vienna,
Austria) was added to the drop, mixed carefully, and allowed
to react for 1 h. Ten min before washing the tips, 5 μL of 1 M
ethanolamine in water was added to the solution in order to
passivate unreacted aldehyde groups. Finally, the tips were
washed three times with PBS buffer and stored in PBS buffer
at 4 °C until use.
Alternatively, coupling via the thiol residue of SH-sgc8c
was done as follows. After APTES silanization as described
before, cantilevers were in a solution of 1 mg NHS-PEG-PDP
dissolved in 500 μL chloroform to which 30 μL TEA were
added, and the cantilevers were allowed to react for 120 min.
Subsequently, they were washed with chloroform and dried
gently by N2 gas. Then the PDP functionalized tips were
immersed in an SH2-sgc8c aptamer solution (10 μM in PBS) for
1 h, washed three times with PBS buffer, and used
immediately or stored in PBS buffer at 4 °C for use within 5 d.
Single molecule force spectroscopy (SMFS)
SMFS experiments were conducted on a PicoPlus 5500 AFM
setup (Keysight, Santa Rosa, USA) equipped with a fluid
chamber to allow measurements in PBS, and an optical
CCD camera to facilitate cantilever alignment with
immobilized T-cells on the substrate. Pulling experiments
were performed in PBS using sgc8c functionalized MSCT
cantilevers with nominal spring constants between 0.01 and
0.03 N m−1 under variation of the pulling velocity to yield
loading rates (i.e., product of pulling velocity and effective
spring constant) ranging from ~400 to 105 pN s−1. The
maximum indentation force was set to 500 pN to avoid any
damage to the cells. At each pulling velocity, 1000 to 2000 force
distance cycles (FDCs) were performed. To ensure
positionindependent results, the position on the cell was shifted by
250 nm μm after 250 FDCs. The delay between approaching
and retraction period (i.e., the hold time) was varied from 0 to
1 s to ensure sufficient contact time for ligand–receptor bond
The spring constant of each cantilever was determined
according to the thermal noise method . Statistical analysis
of all FDCs were done to determine the probability of
aptamer–receptor complex formation (binding probability,
BP), the distribution of detected unbinding forces and lengths,
as well as the effective spring constant (spring constant in the
moment of rupture). The BP is defined as the number of FDCs
exhibiting an unbinding event divided by the total number of
collected FDCs. Each individual rupture force of a single
unbinding event was plotted against its individual corresponding
force loading rate r (determined from the effective spring
constant multiplied by the pulling velocity) and finally merged
into a dynamic force spectra plot. The loading rate-dependent
unbinding forces were evaluated with a maximum likelihood
approach  to fit a statistical model based on the Evans
theory  that allows calculation of the dissociation rate
constant (koff) and the width of the energy barrier xβ based
on the equation fu(r) = (kBT/x)ln[rx/(kBTkoff)], where x is the
separation of the energetic barrier to the equilibrium position,
koff the dissociation constant at zero force, kBT the thermal
energy, and fu(r) the most probable unbinding force at the
loading rate r.
In TREC magnetically coated and ligand functionalized
cantilevers are excited by an alternating magnetic field to oscillate
close to their resonance during scanning. The resonance
frequency of biofunctionalized MACLevers Type VII was
determined by recording a frequency plot, and the actuation
frequency was typically set about 0.5 kHz below the maximum
of the resonance. Then the cantilever was positioned above the
T-cell of interest and slowly approached to avoid damage of
the tip coating and/or the cell. Since the cells appeared very
soft, recognition imaging was only performed on the outer
area of the cells (i.e., the area of the cells that have the higher
distance to the middle of the cell). TREC measurements were
done in PBS buffer at 1 Hz line frequency. The amplitude was
set to a value obtained by force distance cycles, which ensures
sufficient damping of the upper part of the oscillation in case
of molecular (sgc8c–PTK7) interaction, but at the same time
allows remaining in a bound state while measuring above the
recognition site . All other measurement parameters were
identical to the SMFS experiments.
For data evaluation, the threshold of recognition spots was
set to five times the root mean square (rms) of the flattened
recognition image. The spot size for recognition was set to a
typical value of 15 pixels for a 1 × 1 μm scan at 512 pixel/line.
A mask overlay of the determined recognition spots with the
corresponding topographic image was generated using
Gwyddion FreeSPM software (ver. 2.44).
Fig. 1 (A) DNA sequence of the
used sgc8c aptamer. (B) Tip
chemistry. (1) Inert silicon nitride
aminofunctionalized using APTES gas
phase silanization. The
NHS-PEG-PDP (2a) or
NHSPEG-Acetal (2b) is coupled
allowing binding NH2-terminated
(4) DNA aptamers [after
deprotection of the acetal group
(3b)] or SH-terminated sgc8c
aptamers (3a) respectively
Results and discussion
The DNA aptamer sgc8c (Fig. 1A) has been designed to
specifically recognize the receptor PTK7 in the plasma
membrane. It is an important player in extracellular
signaling and highly expressed on the surface of leukemia
cells and especially T-ALL cells. In this study, we used
cultured T-cells of the Jurkat non-Hodgkin’s
lymphoblastic leukemia cell line as an accurate representation
of the native state of PTK7 in lymphoma cells.
For characterization of the specific interaction of
sgc8c with PTK7 on the cell surface at the molecular
level by SMFS, the DNA aptamer was chemically
attached to the apex of the silicon (nitride) tip of the
AFM cantilever via heterobifunctional crosslinkers
(Fig. 1B). After aminofunctionalization (Fig. 1B-1),
depending on the coupling group of the aptamer, which
was either 5′aminofunctionalized (NH2-sgc8c) or 5′
thiolated (SH2-sgc8c), an NHS-PEG-acetal  or an
NHS-PEG-PDP  was covalently bound to the tip,
respectively (Fig. 1B-2a, 2b). Thiolated sgc8c was
coupled without further derivatisation simply through
incubation to the PDP terminus of the tip-bound linker
(Fig. 1B-3a). Binding of NH2-sgc8c required
deprotection of the free acetal end of the linker to produce
an aldehyde residue (Fig. 1B-3b) for reaction with the
amine terminus of the aptamer followed by hydration of
the formed bond after coupling.
T-cells, like all lymphocytes, circulate in the blood stream
without adhesion to blood and lymphatic vessels, and
adhesion and migration through the wall of vessels usually
happens only in response to inflammation. Thus, Cell-Tak was
used as adhesive coating to immobilize T-cells on glass cover
slips for AFM investigation . The cell suspension was
handled carefully to avoid cell lysate of broken cells that
may inactivate Cell-Tak and result in inefficient adherence
of intact T-cells. Next, cells were chemically fixed. This was
necessary since T-cell adhesion as the first step in immune
response is known to start a cascade of changes of the cellular
state that include rolling, cell arrest, strengthening of adhesion
sites, followed by migration . Incubation with 4%
formaldehyde for 60 min was performed, which has been reported
to properly fix cells without loss of functionality of PTK7
receptors on the plasma membrane .
For AFM force spectroscopy measurements, the sgc8c
functionalized cantilever tips were positioned above a fixed
cell using the optics of the AFM setup (Fig. 2A) and
approachretract cycles were executed, and the acting force on the
cantilever was monitored as depicted in Fig. 2B in the form of a
force-distance curve (FDC). In the approach-phase (Fig. 2B
[red curve]) the cantilever starts to experience an increasing
force and bend upward upon contact with the cell. The
nonlinear slope of the first part of bending is a result of partly
compressing of the cell. After reaching a given indentation
force limit of typically 300–400 pN, the retraction phase was
initialized (Fig. 2B [black curve]). In case sgc8c had formed a
complex with PTK7 on the cell surface, further retraction of
the cantilever lead to a pulling force, which appeared as
negative slope in the retract part force due to a downward bending
of the cantilever. When the pulling force exceeded the binding
strength between the aptamer on the tip and PTK7 receptor on
the Jurkat cells, a clear rupture event was observed with a
rupture force corresponding to the sgc8c-PTK7 bond strength.
For statistical analysis, 500–2000 FDCs were recorded
under variation of the location on the cell surface to avoid
measuring artefact. To differentiate these supposed specific
rupture events from nonspecific adhesion, and to avoid measuring
artefact FDCs were collected on different positions on the
cells and the hold time (i.e., the resting or hold time of the
functionalized cantilever on the cell surface) was varied from
0 to 1000 ms, with no observable difference for the number or
appearance of rupture events or measured rupture forces. In a
few cases, double or multiple rupture events were observed.
Such events are either caused by the binding of additional tip
tethered sgc8c molecules to further PTK7 receptors on the cell
or by nonspecific adhesion of the tip with the cell.
Nonetheless, they were not used for the data evaluation since
differentiation of these events cannot be done with sufficient
To prove the specificity of the detected rupture events, the
experiment was performed in the presence of free sgc8c
aptamers in solution (1 μM) to saturate (block) PTK7
and after (red) addition of free ligand. The red curve is normalized to the
relative binding probability. The significance of the proof experiment is
shown in the inset. (D) Probability density functions of sgc8c–T-cell
interactions at different pulling velocities. The most probable unbinding
force increases at higher pulling velocities. The fit of the resulting loading
rate dependence of the rupture force is shown in the inset
receptors on the cell surface. This resulted in a significantly
lowered binding probability. In Fig. 2C the distribution of
measured rupture forces is plotted in the form of a
mathematical probability density function (pdf) before (black) and after
(red) the addition of the blocking is shown for a representative
cell. The red curve is normalized to the relative binding
probability. The existence of a remnant BP may be explained by an
incomplete block of accessible PTK7 receptors on the cell
surface. The averaged binding probability of 38.3 ± 7.48%
before the block was reduced to 10.15 ± 1.93% after addition
of sgc8c to the Jurkat cells (Fig. 2C, inset). An additional,
complementing specificity proof was performed with a tip that
was functionalized with the TDO5 DNA aptamer that
recognizes Ramos cell of the Burkitt’s lymphoma type, but does not
bind to PTK7. The binding probability was only 8.27 ±
The rupture force of a ligand–receptor complex is
dependent on the kinetics of the experiment, in particular the force
ramp of the pulling force acting on the complex during
rupture, and is termed the loading rate [52, 57]. From
extrapolation of the rupture force dependence on the loading rate, the
dissociation rate constant koff and the width of the interaction
potential xβ of the sgc8c-PTK7 interaction on the cell surface
can be calculated. Experimentally, the loading rate is the
product of the pulling speed and the effective spring constant at the
point of rupture. Hence, variation of the loading rate is
achieved by a change of pulling speed of the cantilever. The
effective spring, on the other hand, is determined from a fit of
the slope of the force curve at the point. Datasets were
measured at pulling speeds ranging from 500 to 24000 nm s−1 and
the rupture forces, rupture lengths, and effective spring
constants for each single rupture event were identified. Statistical
analysis at a given pulling speed showed that the increase of
the pulling velocity caused an increase of the rupture forces, as
illustrated by a shift of the maximum of the pdf (Fig. 2D). This
is in good agreement with Evans’ theory for a single energy
barrier in the ligand–receptor interaction potential in the
thermally activated regime . For calculation of koff and xβ
from the loading rate dependence, all individual rupture
events were plotted as a data cloud to account for the influence
of the effective spring constant that varies strongly at a given
pulling velocity as a result of the position-dependent elasticity
of the cell . A fit of the data by a maximum likelihood
approach (as described in more detail in ) yielded a
dissociation constant koff of 5.16 ± 0.19 s−1 with a width of the
energy barrier xβ of 0.65 ± 0.01 Å. The kinetic off rate is
indicative of a slow dissociation of the complex, which is
favorable in terms of an extended interacting time between
the aptamer and the PTK7 in the cell membrane.
In SMFS the functionalized AFM cantilever probes the
surface Bblindly,^ which has an influence on the probability
to form ligand–receptor complexes to measure their rupture.
Whereas the binding probability on a dense layer of isolated
receptors can be as high as 70 to 80%, binding probabilities on
cells range typically below 20% due to a less dense and less
homogenous distribution of receptors and possibly
preferential location in more specified membrane domains . In the
presented study, an uncommonly high average binding
probability of nearly 40% was detected, leading us to investigate
the distribution of PTK7 on the surface of Jurkat T-cells more
closely by TREC.
The method of TREC is illustrated in Fig. 3A; a magnetically
coated cantilever with a quality factor in liquid of ~1 carries an
sgc8c aptamer tethered to the tip via a flexible linker. The tip
oscillates above the cell surface driven by an alternating
magnetic field (MACmode). During lateral scan at a rate of 0.5–1.0 Hz,
sgc8c can bind to PTK7 in the downward swing of the
oscillation, which leads to linker stretching and reduction of the upward
swing. Separation of the bottom part of the oscillation amplitude
that contains the topographic information from the top part that is
only influenced by binding events, a topography image and
recognition map are created from a single scan with lateral
resolution of a few nanometer . To provide for sufficient contact
time that enables ligand–receptor complex formation and in
order to achieve high lateral resolution, surface scans were
performed at 1.5 × 1.5 μm2 scan range (Fig 3B). However, due to
the very soft behavior and high compressibility of the gently
fixed spherical T-cells, TREC imaging on top of a cell turned
out to be more challenging. Hence, all images displayed in Fig. 3
were captured towards the border area of a cell. The topography
revealed a rather smooth surface (Fig. 3B-1, C-1) but a high
number of pronounced dark patches (recognition spots) that
reflect positions of aptamer binding sites on the Jurkat cell
(Fig. 3B-2). Figure 3B-3 shows the overlay of the recognition
events indicated in red with the topography image and show that
PTK7 was evenly distributed over the scan area. In order to
prove that the detected aptamer binding sites were indeed the
locations of PTK7 receptor molecules sgc8c saturated T-cells
were scanned using the identical AFM tips with the same
imaging parameters and settings . Whereas the topography
images showed the same features before (Fig. 3B-1) and after block
(Fig. 3C-1), the recognition signals were nearly completely
abolished (Fig. 3C-2) due to block of the PTK7 receptors by
the added sgc8c DNA aptamers, as illustrated by the cartoon in
Fig. 3C-3, proving that the recognition image in 3B2 shows the
distribution of PTK7 in the cell membrane. The TREC imaging
revealed a homogeneous distribution of PTK7 molecules on the
outer regions of ALL cells with a surface density of 325 ± 12
PTK7 receptors (or small receptor clusters) per μm .
In this study, we demonstrated the specificity of the interaction
of the sgc8c DNA aptamer and PTK7 receptor in the plasma
membrane of intact Jurkat T-cell lymphoma, and measured
Recognition Image Topography Image
Fig. 3 TREC experiments: (A)
Schematic of TREC setup. The
upper part of the oscillation is
used to gain the recognition
image, whereas the lower part is
influenced by the sample
topography. Topography (B1) and
recognition (B2) image on a
Tcell membrane using sgc8c
functionalized tips. A superposition of
topography and recognition is
shown at image (B3). After
addition of free aptamers the
topography (C1) remains unchanged,
whereas the recognition spots
(C2) is completely abolished as a
result of blocked PTK7 receptors
(illustrated in C3). Scale bar for
all AFM images is 500 nm
rupture forces of the ligand–receptor complex under different
loading rates using SMFS. The kinetic off rate of the studied
system (koff = 5.16 s−1) indicates slow dissociation of the
complex. Furthermore, we demonstrated the possibility of
performing recognition imaging experiments on T-cells for
the first time. TREC represents a powerful imaging tool in
which topography and recognition of specific biological
molecules are simultaneously mapped. Our results show that
aptamers covalently coupled to an AFM probe recognize
specific receptors at nanometer lateral resolution and with high
specificity. More importantly, we were able to visualize and
quantify the distribution of PTK7 receptors in the cell
membrane, showing a high density and homogenous lateral
distribution in ALL-cells, making it an ideal target. This study
provides new insight into the mode of the action of the sgc8c
DNA aptamers as a diagnostic and targeting agent for acute
lymphoblastic leukemia. As the continuing advancements of
the cell SELEX technique  yield a growing library of
aptamers for various cancer markers, they may prove
particularly useful for identification and capturing of circulating
tumor cells (CTC) . Moreover, our study presents validation
of the method of AFM biosensing for the detection of cancer
markers and reveals potential for future clinical diagnostics. In
combination with traditional histology-based analysis of
biopsies, the method could reduce tumor misclassification .
Also, with the expression level of tumor markers being related
to the clinical stage of the disease, determination of the marker
density on the cell surface by TREC might allow for
assessment of disease progression as well as the efficacy of cancer
Acknowledgements Open access funding provided by Johannes
Kepler University Linz. This work was supported by OEAD, by
Agency for Promotion Research and Development (project no.
APVV14-0267 and SK-AT-2015-0004), by grant agency VEGA (project no.
2/0055/14), and EU Horizon 2020 Marie Sklodowska-Curie Grant
656842. The authors are grateful for support by Dr. Christian Rankl in
data evaluation and with the evaluation software.
Compliance with ethical standards
Conflict of interest The authors declare no conflict of interest.
This article does not contain any research with human participants or
Open Access This article is distributed under the terms of the Creative
C o m m o n s A t t r i b u t i o n 4 . 0 I n t e r n a t i o n a l L i c e n s e ( h t t p : / /
creativecommons.org/licenses/by/4.0/), which permits unrestricted use,
distribution, and reproduction in any medium, provided you give
appropriate credit to the original author(s) and the source, provide a link to the
Creative Commons license, and indicate if changes were made.
Michael Leitner received his
PhD in Biophysics in 2012 from
the Johannes Kepler University
Linz, Austria. He is a biophysicist
and medical technique engineer
with more than 10 years of
experience in bio-AFM, topography,
and recognition imaging and
single molecule force spectroscopy.
A l e x a n d r a P o t u r n a y o v a
defended her PhD thesis in
2012 at the Institute of Animal
Biochemistry and Genetics of the
Slovak Academy of Sciences,
Ivanka pri Dunaji, Slovakia. She
is a research fellow at this institute
and partially at the Faculty of
M a t h e m a t i c s , P h y s i c s , a n d
I n f o r m a t i c s , C o m e n i u s
University in Bratislava. Her
scientific interests include
masssensitive biosensors and surface
characterization by AFM.
Constanze Lamprecht received
her PhD in Biophysics in 2010
f r o m t h e J o h a n n e s K e p l e r
University Linz (JKU) (Austria)
followed by post-doctoral
posit i o n s a t t h e U n i v e r s i t y o f
Waterloo (Canada) and the
University of Kiel (Germany).
She is currently affiliated again
with the Institute of Biophysics
at JKU where she uses AFM and
fluorescence microscopy to study
cell transformations that lead to
the initiation and progression of
Sabine Weich successfully
finished her Master’s degree in
B i o t e c h n o l o g y a n d
Environmental Technology in
2 0 1 3 a t t h e U n i v e r s i t y o f
Applied Sciences, Wels, Austria,
and performed AFM-based
molecular recognition studies at the
Johannes Kepler University,
Maja Snejdarkova defended her
PhD thesis in 1980 at Masaryk
U n i v e r s i t y, B r n o , C z e c h
Republik. She was a research
officer at the Institute of Animal
Biochemistry and Genetics of the
Slovak Academy of Sciences,
Ivanka pri Dunaji, Slovakia. In
2016 she retired, but continued
working in Professor T. Hianik’s
laboratory. Her scientific interests
include mass-sensitive and
Ivana Karpisova is an external
PhD student at the Faculty of
M a t h e m a t i c s , P h y s i c s , a n d
I n f o r m a t i c s , C o m e n i u s
University, Bratislava, Slovakia,
working under supervision of
Professor T. Hianik. Her work is
focused on mass-sensitive
biosensors and AFM. Since 2016 she
has been working as associate
s a f e t y r e v i e w s p e c i a l i s t a t
Andreas Ebner received his PhD
in Biophysics in 2007. Since 2016
he has been Associate Professor at
the Institute of Biophysics and
Deputy Head of the Department
o f A p p l i e d E x p e r i m e n t a l
Biophysics at the Johannes
Kepler University Linz, Austria.
His research expertise includes
biosensing atomic force
microscopy and spectroscopy, and single
molecule sensor tip chemistry.
1. Jemal A , Siegel R , Ward E , Murray T , Xu J , Thun MJ . Cancer statistics, 2007 . CA Cancer J Clin . 2007 ; 57 : 43 - 66 .
2. Kawasaki ES , Clark SS , Coyne MY , Smith SD , Champlin R , Witte ON , et al. Diagnosis of chronic myeloid and acute lymphocytic leukemias by detection of leukemia-specific messenger-RNA sequences amplified in vitro . Proc Natl Acad Sci U S A . 1988 ; 85 : 5698 - 702 .
3. Ramaswamy S , Tamayo P , Rifkin R , Mukherjee S , Yeang CH , Angelo M , et al. Multiclass cancer diagnosis using tumor gene expression signatures . Proc Natl Acad Sci U S A . 2001 ; 98 : 15149 - 54 .
4. Mason J , Griffiths M. Molecular diagnosis of leukemia . Expert Rev Molec Diag . 2012 ; 12 : 511 - 26 .
5. Craig FE , Foon KA . Flow cytometric immunophenotyping for hematologic neoplasms . Blood . 2008 ; 111 : 3941 - 67 .
6. Eckert C , Biondi A , Seeger K , Cazzaniga G , Hartmann R , Beyermann B , et al. Prognostic value of minimal residual disease in relapsed childhood acute lymphoblastic leukemia . Lancet . 2001 ; 358 : 1239 - 41 .
7. Haferlach T , Kern W , Schnittger S , Schoch C. Modern diagnostics in acute leukemias . Crit Rev Oncol Hematol . 2005 ; 56 : 223 - 34 .
8. Wu X , Chen J , Wu M , Zhao JX . Aptamers: active targeting ligands for cancer diagnosis and therapy . Theranostics . 2015 ; 5 : 322 - 44 .
9. Zhu GZ , Ye M , Donovan MJ , Song E , Zhao Z , Tan W. Nucleic acid aptamers: an emerging frontier in cancer therapy . Chem Commun . 2012 ; 48 : 10472 - 80 .
10. Radom F , Jurek PM , Mazurek MP , Otlewski J , Jeleń F. Aptamers : molecules of great potential . Biotech Adv . 2013 ; 31 : 1260 - 74 .
11. Zhou JH , Rossi JJ . Cell type-specific, aptamer-functionalized agents for targeted disease therapy . Mol Therapy-Nucleic Acids . 2014 ; 3 , e169.
12. Zhu GZ , Niu G , Chen X. Aptamer-drug conjugates. Bioconjug Chem . 2015 ; 26 : 2186 - 97 .
13. Yang ML , Jiang GH , Li W , Qiu K , Zhang M , Carter CM , et al. Developing aptamer probes for acute myelogenous leukemia detection and surface protein biomarker discovery . J Hematol Oncol . 2014 ; 7 : 5 .
14. Shangguan D , Li Y , Tang Z , Cao ZC , Chen HW , Mallikaratchy P , et al. Aptamers evolved from live cells as effective molecular probes for cancer study . Proc Natl Acad Sci U S A . 2006 ; 103 : 11838 - 43 .
15. Shangguan D , Tang ZW , Mallikaratchy P , Xiao Z , Tan W. Optimization and modifications of aptamers selected from live cancer cell lines . Chem Biochem . 2007 ; 8 : 603 - 6 .
16. Peradziryi H , Tolwinski NS , Borchers A. The many roles of PTK7: a versatile regulator of cell-cell communication . Arch Biochem Biophys . 2012 ; 524 : 71 - 6 .
17. Gartner S , Gunesch A , Knyazeva T , Wolf P , Högel B , Eiermann W , et al. PTK 7 is a transforming gene and prognostic marker for breast cancer and nodal metastasis involvement . PLoS One . 2014 ; 9 , e84472.
18. Kim JH , Kwon J , Lee HW , Kang MC , Yoon HJ , Lee ST , et al. Protein tyrosine kinase 7 plays a tumor suppressor role by inhibiting ERK and AKT phosphorylation in lung cancer . Oncol Rep . 2014 ; 31 : 2708 - 12 .
19. Zhang HT , Wang AD , Qi S , Cheng S , Yao B , Xu Y. Protein tyrosine kinase 7 (PTK7) as a predictor of lymph node metastases and a novel prognostic biomarker in patients with prostate cancer . Int J Mol Sci . 2014 ; 15 : 11665 - 77 .
20. Huang YF , Shangguan DH , Liu H , Phillips JA , Zhang X , Chen Y , et al. Molecular assembly of an aptamer-drug conjugate for targeted drug delivery to tumor cells . Chem Biochem . 2009 ; 10 : 862 - 8 .
21. Taghdisi SM , Abnous K , Mosaffa F , Behravan J. Targeted delivery of daunorubicin to T-cell acute lymphoblastic leukemia by aptamer . J Drug Target . 2010 ; 18 : 277 - 81 .
22. Zhu GZ , Zheng J , Song E , Donovan M , Zhang K , Liu C , et al. Self-assembled, aptamer-tethered DNA nanotrains for targeted transport of molecular drugs in cancer theranostics . Proc Natl Acad Sci U S A . 2013 ; 110 : 7998 - 8003 .
23. Huang YF , Sefah K , Bamrungsap S , Chang HT , Tan W. Selective photothermal therapy for mixed cancer cells using aptamer-conjugated nanorods . Langmuir . 2008 ; 24 : 11860 - 5 .
24. Wang J , You MX , Zhu G , Shukoor MI , Chen Z , Zhao Z , et al. Photosensitizer-gold nanorod composite for targeted multimodal therapy . Small . 2013 ; 9 : 3678 - 84 .
25. Kang HZ , O'Donoghue MB , Liu H , Tan W. A liposome-based nanostructure for aptamer directed delivery . Chem Commun . 2010 ; 46 : 249 - 51 .
26. Shi H , He XX , Wang K , Wu X , Ye X , Guo Q , et al. Activatable aptamer probe for contrast-enhanced in vivo cancer imaging based on cell membrane protein-triggered conformation alteration . Proc Natl Acad Sci U S A . 2011 ; 108 : 3900 - 5 .
27. Jacobson O , Weiss ID , Wang L , Wang Z , Yang X , Dewhurst A , et al. F-18-labeled single-stranded DNA aptamer for PET imaging of protein tyrosine kinase-7 expression . J Nuclear Med . 2015 ; 56 : 1780 - 5 .
28. Li H , Hu HT , Zhao Y , Chen X , Li W , Qiang W , et al. Multifunctional aptamer-silver conjugates as theragnostic agents for specific cancer cell therapy and fluorescenceenhanced cell imaging . Anal Chem . 2015 ; 87 : 3736 - 45 .
29. Zheng FY , Cheng Y , Wang J , Lu J , Zhang B , Zhao Y , et al. Aptamer-functionalized barcode particles for the capture and detection of multiple types of circulating tumor cells . Adv Mater . 2014 ; 26 : 7333 - 8 .
30. O'Donoghue MB , Shi XL , Fang X , Tan W. Single-molecule atomic force microscopy on live cells compares aptamer and antibody rupture forces . Anal Bioanal Chem . 2012 ; 402 : 3205 - 9 .
31. Lamprecht C , Hinterdorfer P , Ebner A. Applications of biosensing atomic force microscopy in monitoring drug and nanoparticle delivery . Expert Opin Drug Del . 2014 ; 11 : 1237 - 53 .
32. Wang N , Liu NQ , Hao J , Bai X , Li H , Zhang Z , et al. Single molecular recognition force spectroscopy study of a DNA aptamer with the target epithelial cell adhesion molecule . Analyst . 2015 ; 140 : 6226 - 9 .
33. Beaussart A , Abellan-Flos M , El-Kirat-Chatel S , Stéphane P , Vincent SP , Dufrêne YF . Force nanoscopy as a versatile platform for quantifying the activity of antiadhesion compounds targeting bacterial pathogens . Nano Lett . 2016 ; 16 : 1299 - 307 .
34. Evans E. Energy landscapes of biomolecular adhesion and receptor anchoring at interfaces explored with dynamic force spectroscopy . Faraday Discus . 1998 ; 111 : 1 - 16 .
35. Friddle RW , Noy A , De Yoreo JJ . Interpreting the widespread nonlinear force spectra of intermolecular bonds . Proc Natl Acad Sci U S A . 2012 ; 109 : 13573 - 8 .
36. Alsteens D , Pfreundschuh M , Zhang CH , Spoerri PM , Coughlin SR , Kobilka BK , et al. Imaging G protein-coupled receptors while quantifying their ligand-binding free-energy landscape . Nat Methods . 2015 ; 12 : 845 - 51 .
37. Dufrene YF , Martinez-Martin D , Medalsy I , Alsteens D , Muller DJ . Multiparametric imaging of biological systems by force-distance curve-based AFM . Nat Methods . 2013 ; 10 : 847 - 54 .
38. Li Q , Zhang T , Pan YG , Ciacchi LC , Xu BQ , Wei G . AFM-based force spectroscopy for bioimaging and biosensing . RSC Adv . 2016 ; 6 : 12893 - 912 .
39. Stroh C , Wang H , Bash R , Ashcroft B , Nelson J , Gruber H , et al. Single-molecule recognition imaging microscopy . Proc Natl Acad Sci U S A . 2004 ; 101 : 12503 - 7 .
40. Stroh CM , Ebner A , Geretschläger M , Freudenthaler G , Kienberger F , Kamruzzahan AS , et al. Simultaneous topography and recognition imaging using force microscopy . Biophys J . 2004 ; 87 : 1981 - 90 .
41. Leitner M , Mitchell N , Kastner M , Schlapak R , Gruber HJ , Hinterdorfer P , et al. Single-molecule AFM characterization of individual chemically tagged DNA tetrahedra . ACS Nano . 2011 ; 5 : 7048 - 54 .
42. Leitner M , Stock LG , Traxler L , Leclercq L , Bonazza K , Friedbacher G , et al. Mapping molecular adhesion sites inside SMIL coated capillaries using atomic force microscopy recognition imaging . Anal Chim Acta . 2016 ; 930 : 39 - 48 .
43. Tang J , Ebner A , Badelt-Lichtblau H , Völlenkle C , Rankl C , Kraxberger B , et al. Recognition imaging and highly ordered molecular templating of bacterial S-layer nanoarrays containing affinity-tags . Nano Lett . 2008 ; 8 : 4312 - 9 .
44. Ebner A , Nikova D , Lange T , Häberle J , Falk S , Dübbers A , et al. Determination of CFTR densities in erythrocyte plasma membranes using recognition imaging . Nanotechnology . 2008 ; 19 : 384017 .
45. Chtcheglova LA , Atalar F , Ozbek U , Wildling L , Ebner A , Hinterdorfer P. Localization of the ergtoxin-1 receptors on the voltage sensing domain of hERG K+ channel by AFM recognition imaging . Pflugers Arch - Eur J Physiol . 2008 ; 456 : 247 - 54 .
46. Duman M , Pfleger M , Zhu R , Rankl C , Chtcheglova LA , Neundlinger I , et al. Improved localization of cellular membrane receptors using combined fluorescence microscopy and simultaneous topography and recognition imaging . Nanotechnology . 2010 ; 21 : 115504 .
47. Ebner A , Hinterdorfer P , Gruber HJ . Comparison of different aminofunctionalization strategies for attachment of single antibodies to AFM cantilevers . Ultramicroscopy . 2007 ; 107 : 922 - 7 .
48. Wildling L , Unterauer B , Zhu R , Rupprecht A , Haselgrübler T , Rankl C , et al. Linking of sensor molecules with amino groups to amino-functionalized AFM tips . Bioconj Chem . 2011 ; 22 : 1239 - 48 .
49. Kamruzzahan ASM , Ebner A , Wildling L , Kienberger F , Riener CK , Hahn CD , et al. Antibody linking to atomic force microscope tips via disulfide bond formation . Bioconjug Chem . 2006 ; 17 : 1473 - 81 .
50. Butt HJ , Jaschke M. Calculation of thermal noise in atomic force microscopy . Nanotechnology . 1995 ; 6 : 1 - 7 .
51. Neundlinger I , Puntheeranurak T , Wildling L , Rankl C , Wang LX , Gruber HJ , et al. Forces and dynamics of glucose and inhibitor binding to sodium glucose co-transporter SGLT1 studied by single molecule force spectroscopy . J Biol Chem . 2014 ; 289 : 21673 - 83 .
52. Evans E , Ritchie K. Dynamic strength of molecular adhesion bonds . Biophys J . 1997 ; 72 : 1541 - 55 .
53. Preiner J , Ebner A , Chtcheglova L , Zhu R , Hinterdorfer P. Simultaneous topography and recognition imaging: physical aspects and optimal imaging conditions . Nanotechnology . 2009 ; 20 : 215103 .
54. Jena B. Fusion pore or porosome: structure and dynamics . J Endocrinol . 2003 ; 176 : 169 - 74 .
55. Schmidt S , Moser M , Sperandio M. The molecular basis of leukocyte recruitment and its deficiencies . Mol Immunol . 2013 ; 55 : 49 - 58 .
56. Thavarajah R , Mudimbaimannar VK , Elizabeth J , Rao UK , Ranganathan K. Chemical and physical basics of routine formaldehyde fixation . J Oral Maxillofacial Pathol . 2012 ; 16 : 400 - 5 .
57. Baumgartner W , Hinterdorfer P , Ness W , Raab A , Vestweber D , Schindle H , et al. Cadherin interaction probed by atomic force microscopy . Proc Natl Acad Sci U S A . 2000 ; 97 : 4005 - 10 .
58. Lamprecht C , Plochberger B , Ruprecht V , Wieser S , Rankl C , Heister E , et al. A single-molecule approach to explore binding, uptake, and transport of cancer cell targeting nanotubes . Nanotechnology . 2014 ; 25 : 125704 .
59. Hu YS , Cang H , Lillemeier BF. Super-resolution imaging reveals nanometer- and micrometer-scale spatial distributions of T-cell receptors in lymph nodes . Proc Natl Acad Sci U S A . 2016 ; 113 : 7201 - 6 .
60. Zhu X , Lu PY , Rosato RR , Tan W , Zu Y. Oligonucleotide aptamers: new tools for targeted cancer therapy . Molec Therapy - Nucleic Acids . 2014 ; 3 , e182.
61. Dickey DD , Giangrande PH . Oligonucleotide aptamers: A next-generation technology for the capture and detection of circulating tumor cells . Methods . 2016 ; 97 : 94 - 103 .
62. Spence T , De Souza R , Dou Y , Stapleton S , Reilly RM , Allen C. Integration of imaging into clinical practice to assess the delivery and performance of macromolecular and nanotechnology-based oncology therapies . J Control Release . 2015 ; 219 : 295 - 312 .