ARTICLE)

Improved Single Molecule Force
Spectroscopy Using Micromachined
Cantilevers
Matthew S. Bull,†,‡, Ruby May A. Sullan,†,# Hongbin Li,§ and Thomas T. Perkins†,^,*JILA, National Institute of Standards and Technology and University of Colorado, Boulder, Colorado 80309, United States, ‡Department of Physics, University ofColorado, Boulder, Colorado 80309, United States, §Department of Chemistry, University of British Columbia, Vancouver, British Columbia V6T 1Z1, Canada, and ^Departmentof Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, Colorado 80309, United States. Present address: Department of Applied Physics, Stanford
University, Stanford, California 94305, United States. #Present address: Institute of Life Sciences, Université Catholique de Louvain, B-1348 Louvain-la-Neuve, Belgium.
)†

ABSTRACT Enhancing the short-term force precision of atomic

force microscopy (AFM) while maintaining excellent long-term force
stability would result in improved performance across multiple AFM
modalities, including single molecule force spectroscopy (SMFS). SMFS
is a powerful method to probe the nanometer-scale dynamics andenergetics of biomolecules (DNA, RNA, and proteins). The folding andunfolding rates of such macromolecules are sensitive to sub-pN
changes in force. Recently, we demonstrated sub-pN stability over abroad bandwidth (Δf = 0.0116 Hz) by removing the gold coatingfrom a 100 μm long cantilever. However, this stability came at the cost of increased short-term force noise, decreased temporal response, and poor sensitivity.
Here, we avoided these compromises while retaining excellent force stability by modifying a short (L = 40 μm) cantilever with a focused ion beam. Our process
led to a ∼10-fold reduction in both a cantilever's stiﬀness and its hydrodynamic drag near a surface. We also preserved the beneﬁts of a highly reﬂective
cantilever while mitigating gold-coating induced long-term drift. As a result, we extended AFM's sub-pN bandwidth by a factor of ∼50 to span ﬁve decades of
bandwidth (Δf ≈ 0.011000 Hz). Measurements of mechanically stretching individual proteins showed improved force precision coupled with state-of-the-art
force stability and no signiﬁcant loss in temporal resolution compared to the stiﬀer, unmodiﬁed cantilever. Finally, these cantilevers were robust and werereused for SFMS over multiple days. Hence, we expect these responsive, yet stable, cantilevers to broadly beneﬁt diverse AFM-based studies.
KEYWORDS: AFM . atomic force microscopy . protein folding . SMFS . focused ion beam milling . cantilever dynamics .
e-beam induced depositionA

tomic force microscopy (AFM)1 is a
powerful tool in nanoscience that is
having an increasing impact in biology.210 AFM oﬀers subnanometer imaging7,1114 in conjunction with mechanicalprobing of molecules10,1517 and cells.18,19
The primary measurement in AFM is force(F), derived from measuring cantilever deﬂection. Like any measurement platform,
AFM would beneﬁt from better precisionon short time scales while maintaining excellent stability over long periods. Better
short-term force precision improves all
AFM applications. Likewise, long-term
force stability beneﬁts multiple AFM modalities, such as imaging20,21 and single
molecule force spectroscopy (SMFS).15 In
SMFS, force stability is critical since theequilibrium between folded and unfolded
BULL ET AL.

states of biomolecules (proteins, RNA,and DNA) is sensitive to sub-pN changes
in F.15,22
The path toward improved short-term
force precision is well established:23 reduce
the hydrodynamic drag (β) of the cantilever.
This improvement is a consequence ofthe ﬂuctuationdissipation theorem ΔF =(4kBTΔfβ)1/2, where ΔF is the force precision, kBT is the thermal energy, and Δf is the
bandwidth of the measurement. The primary way to reduce β is to decrease thecantilever length (L), which has led to shorter, albeit stiﬀer, cantilevers.23 Stiﬀer cantilevers do not adversely aﬀect force precision as long as the measured motion of thecantilever is dominated by Brownian motion(an assumption of the ﬂuctuationdissipation
theorem).
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Received for review February 21, 2014
and accepted March 26, 2014.
Published online
10.1021/nn5010588
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ARTICLEIn contrast, long-term stability is limited by instrumental rather than fundamental issues. Recently, weachieved sub-pN force stability over a 100-s period(Δf = 0.0116 Hz) in liquids by removing the gold
coating of a soft (k = 6 pN/nm) cantilever. Positional
precision at higher frequencies (∼1 kHz) remained
unchanged despite a ∼10-fold reduction in reﬂected
light.24 The enhanced stability aﬀorded by an uncoated cantilever revealed low-frequency instrumental
drift in the optical lever arm. This drift was similar inmagnitude and spectral distribution on a state-of-theart commercial AFM (Cypher, Asylum Research) and acustom-built ultrastable AFM.25 The drift added a ﬁxed
amount of additional positional noise per unit bandwidth, which in turn degraded the performance ofstiﬀer cantilevers more than softer ones (Figure 1). As aresult, cantilevers that are soft and long (Figure 1a)outperformed short yet stiﬀ ones (Figure 1b) on surprisingly short time scales (∼25 ms) when using two
popular cantilevers, the long BioLever [k = 6 pN/nm;
L = 100 μm (Olympus)] and the BioLever Mini [k =100 pN/nm; L = 40 μm].21
An additional goal in SMFS is detecting small, shortlived folding intermediates.26,27 Such detection is enhanced by improved short-term force precision andfast response of the cantilever to an abrupt change in F.
Unfortunately, the long BioLevers that provide the bestforce stability suﬀer from relatively poor temporal
resolution due to their increased β and decreased k.
This reduced temporal resolution, coupled with decreased short-term force precision, hinders their application in such SMFS studies.
Thus, at the moment, choosing the appropriatecantilever for a particular application requires a compromise. This choice arises from the basic scaling
relation governing cantilever stiﬀness: k  wT3/L3,
where w is the width of a rectangular cantilever, andT is its thickness. Reductions in β arising from decreased L necessarily lead to increased k. As a result,a user can have a stiﬀ, low-force noise cantilever at thecost of reduced long-term stability. Alternatively, one
can get state-of-the-art long-term force stability, butwith increased force noise per unit bandwidth andreduced temporal resolution. For applications that
require high reﬂectivity, gold-coated cantilevers leadto particularly poor long-term force stability.
We avoided these compromises by modifying ashort (L = 40 μm) commercial cantilever using a
focused ion-beam (FIB) (Figure 1c). Micromachiningof cantilevers with a FIB has led to signiﬁcant drops inboth k and β.28,29 However, these beneﬁts have not yet
been exploited in biophysical and nanoscience research applications using the traditional optical leverarm detection available on commercial AFMs.
In this paper, we developed an eﬃcient protocol forenhancing the spatial-temporal measurement limits ofa widely used commercial cantilever (BioLever Mini)

Figure 1. Micromachined cantilevers show improved performance. SEM images of (a) an uncoated long BioLever (L =100 μm), (b) an uncoated BioLever Mini (L = 40 μm), and (c) a
FIB-modiﬁed BioLever Mini that has a transparent glass
capping layer protecting an underlying gold patch at theend of the cantilever. Each cantilever's measured spring
constant is indicated. (d) The averaged force power spectral
density (PSD) as a function of frequency of each cantilever inliquid is shown when the cantilever was 50 nm over thesurface. The color of the curve is associated with the line
immediately below each cantilever's image in (ac). (e)Integrated force noise for each cantilever shown as afunction of frequency. The sub-pN regime is shaded orange.
The gold patch in panel (c) is false colored for visual clarity.

(Figure 2) and explored its application in precision
protein-unfolding experiments. Removing a large section of the cantilever led to a 10-fold reduction in thehydrodynamic drag, improving short-term force precision. We simultaneously achieved a 10-fold reduction
in stiﬀness by thinning the remaining beams, facilitating excellent long-term force stability. A transparent
capping layer patterned at the end of the cantileverallowed us to retain a gold-coated cantilever's highreﬂectivity while removing the gold from the rest of thecantilever for improved force stability. As a result, we
extended the AFM's sub-pN bandwidth by a factor of∼50 to span a total of ﬁve decades of bandwidth (Δf ≈0.011000 Hz). Our FIB-modiﬁed cantilevers could bereused in SMFS assays over multiple days, increasing
their cost eﬀectiveness. In protein-based SMFS assays,
we demonstrated improved short-term force precisioncoupled with state-of-the-art force stability. Moreover,these soft, but short, cantilevers suﬀered no signiﬁcant
loss in temporal resolution when measuring abrupt
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transitions due to protein unfolding, as compared tothe stiﬀer unmodiﬁed cantilever. Overall, this combination of improved force precision without loss of
temporal resolution, force stability, or sensitivity opens
the door to many exciting AFM-based studies of shortlived nanoscale events, including single-molecule
studies of protein folding over extended periods(100 s), a duration 25-fold longer than previous equilibrium AFM studies.15
RESULTS AND DISCUSSION
Our goal was to improve the performance of AFM ina broad range of biophysical and nanoscience applications while retaining a high-level of usability at anaﬀordable cost. Ease of use was achieved, in part, bypreserving a small gold-coated region on the back of
the cantilever while minimizing drift induced by thegold coating. The long-term force stability, which was
limited by instrumental drift in the optical lever arm,
was achieved by lowering k. Short-term force precision was limited by the hydrodynamic drag, a consequence of the ﬂuctuationdissipation theorem. Thus,
we sought to simultaneously reduce k and β while
preserving the high reﬂectivity in an eﬃcient process.
Highly Reflective Micromachined Cantilevers. In prior
work, we showed that removing a cantilever's gold coating enabled sub-pN force precision over a broad bandwidth, but also led to a 10-fold loss in reflectivity.24 The
decreased optical signal did not adversely affect positional precision on soft cantilevers (k = 6 pN/nm) at
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Figure 2. A four-step protocol for micromachined cantilevers. (a) A commercially available cantilever (Biolever Mini) is
selected for modiﬁcation. (b) A glass-like capping layer isdeposited via e-beam-induced deposition. (c) A focused ion
beam (FIB) cuts out the central region of the cantilever. (d)Using a defocused beam, the narrow cantilever supports arethinned. (e) Two 40 s wet etches remove all of the unprotected gold and chromium to improve low-frequency
performance.

∼1 kHz on a state-of-the-art commercial AFM, but did
decrease the usability of uncoated cantilevers. For example, they showed more pronounced interferenceinduced oscillations in the cantilever signal,21 particularly on gold-coated substrates commonly used insingle-molecule30,31 and nanoscience applications.32,33
Reduced reflectivity is also expected to adversely impact
performance when using stiffer cantilevers at higher
frequencies (>10 kHz) and on older instruments.
A cantilever is most sensitive to the adverse eﬀects
of gold at the region of highest curvature, the junction
between the base of the cantilever and the chip on
which it is mounted. Previously, researchers havecreated spatial patterning of gold near the end of a
cantilever by FIB milling oﬀ a cantilever's gold
coating.34 Similarly, we fully removed the gold andunderlying chromium layer using an FIB. These cantilevers exhibited improved low-frequency performance.
However, we consistently observed better low-frequency
performance by using a wet chemical etch.
To preserve reﬂectivity while also achieving excellent long-term stability, we developed an e-beampatterned capping layer to preserve a small, welldeﬁned section of gold on the back side of thecantilever. This capping layer protected the cantilever's
gold coating during a subsequent wet etch that removed both the gold and chromium from the rest of
the cantilever. Importantly, the capping layer was
made from tetraethyl orthosilicate (TEOS), a glass-like
substance that is optically transparent (see Methods).
As a result, the 3040 nm thick TEOS layer did notadversely aﬀect optical-lever-arm detection. Fully
FIB-modiﬁed cantilevers, as shown in Figure 1c, retained excellent sensitivity (∼1520 nm/V) whenusing the Cypher's standard (not small) spot size
module in comparison to an uncoated BioLever Mini(∼130 nm/V) and showed a minimal (25%) loss in the
optical signal. If necessary, larger reﬂective regions canbe fabricated by extending the TEOS patch over the fullarea of the end of the cantilever.
Reducing Cantilever Stiffness. We initially reduced k byremoving a larger rectangular area (20  14 μm2) fromthe base of a BioLever Mini. The starting width of thecantilever was ∼16 μm, and our modification left two
1 μm wide supporting beams (Figure 1c). These supports were positioned at the edge of cantilever toretain torsional stiffness. Finite-element modelingand the linear scaling of k with w suggested an 8-fold
reduction in w should lead to an 8-fold reduction in k.
Experimentally, we measured a ∼6-fold reduction.
Additional reduction in k can be made by further
narrowing of the support beams; we have made beams
as narrow as 600 nm, but this was technically demanding. Instead, we focused on reducing the thicknessof the cantilever, because k ∼ T3. A defocused FIB
scanned along the long axis of the beam reduced
the thickness from ∼170 to ∼120 nm. This thinning

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and, hence, better short-term force precision. The
degree of improvement can be quantiﬁed by taking
the ratio of the ﬂat sections of the PSD for the twocantilevers. This analysis shows a 3.5-fold improvement, despite the modiﬁed cantilever being 14-foldless stiﬀ.
The kinetics of folding and unfolding of molecules
are sensitive to the total applied force. Thus, a key
metric is the force stability over the full duration of theexperiment. Current state-of-the-art AFM experiments
measure the equilibrium folding and unfolding of aprotein over a few seconds.15 In contrast, equilibrium
assays with dual-beam optical-trapping experiments
may last tens to hundreds of seconds.37,38 To span bothcurrent and future AFM experiments, we calculated theintegrated force noise over a 100-s period, similar toour prior work.24 The beneﬁts of micromachined cantilevers are immediately obvious (Figure 1e). Their
reduced k allows them to have the long-term force
stability equivalent to a long BioLever. Yet, their reduced β allows the integrated force noise remained at
sub-pN levels to 930 Hz, a signiﬁcantly higher f thanthe long BioLever (16 Hz). Hence, the FIB-modiﬁed
cantilevers exhibited a ∼50-fold increase in subpN bandwidth to span ﬁve decades of bandwidth(Δf ≈ 0.011000 Hz).
Comparison with Highly Reflective Cantilevers. For applications and instruments that require high reflectivity,
it is useful to compare our FIB-modified cantilevers
with two traditional gold-coated cantilevers (the long
BioLever and the BioLever Mini). A useful metric tocharacterize performance over varying averaging
times is the Allan variance39 (see Methods). The Allan
variance at a given averaging time represents theaveraged force noise over that time interval. More
broadly, a plot of the Allan variance is another metricto characterize the crossover between thermally limited and instrumentation-limited performance.40 We
computed the Allan variance for isolated cantilevers
held 50-nm over the surface (Figure 3). The Allan
variance initially increases, suggesting worse performance at longer times. However, this is an artifact due
to correlated motion of the cantilever at short timescales;21 this portion of the curve is de-emphasized byplotting it in gray. Over longer averaging times, theforce noise drops with a slope consistent with averaging Brownian motion. At sufficiently long times, theAllan variance for all three cantilevers deviates fromthis slope. At this point, instrumental noise starts tocontribute and eventually dominates the average force
noise on the longest time scalesComparison of the Allan variance for the threehighly reﬂective cantilevers immediately shows the
beneﬁt of the micromachined cantilevers. Over all timescales investigated (0.0000550 s), the modiﬁed cantilever outperforms both commercial cantilevers. This
performance improvement is particularly pronounced
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resulted in a total reduction in k by a factor of ∼10[9.4 ( 0.9 (N = 10)]. For the particular cantilever shownin Figure 1c, the unmodified k was initially 54 pN/nm
and reduced to 4.7 pN/nm, a 12-fold reduction. Further
thinning of the cantilever is possible, though it resulted
in a rapidly increasing fraction of cantilevers breaking during such thinning. Overall, this micromachining process generated the desired short, but soft,
cantilevers.
Characterizing the Performance of Micromachined Cantilevers. The dominant noise sources in AFM-based biophysical measurements (i.e., Brownian motion, instrumental noise in the optical lever arm, and metallizationinduced drift of the cantilever) also affect a simpler
measurement: the zero force position (z0) of the cantilever. Our initial characterization of the FIB-modified
cantilevers focused on this simpler measurement. Moreover, we performed this characterization 50 nm over thesurface, since most AFM-based assays are done in close
proximity to a surface. This proximity increases a cantilever's hydrodynamic damping, a phenomenon called
squeezed-film damping in AFM28 and conceptuallysimilar to Faxen's law in optical-trapping studies.35
Increased damping near a surface degrades short-term
force precision.36
Characterizing cantilever motion as a function of
frequency (f) allows for easy identiﬁcation of two
regimes: thermal- and instrumentation-limited performance. The force power spectral density (PSD) was
calculated from a set of ﬁve 100 s records for eachcantilever (Figure 1d) (Methods). The thermally limited
regime is at higher f, where the PSD is ﬂat as a functionof f and then starts to fall oﬀ at a characteristic roll-oﬀ
frequency analogous to the PSD of optically trapped
beads.35 The instrumentation-limited regime is seen atlower f, where the PSD increases as f decreases. The
crossover between these two regimes diﬀers for thethree cantilevers studied.
The two uncoated commercial cantilevers show
better performance in diﬀerent regimes.24 The
BioLever Mini outperformed the long BioLever in thethermally limited regime (higher f) because of its lower
hydrodynamic drag relative to the long BioLever. On
the other hand, the long BioLever outperformed the
BioLever Mini in the instrumentation-limited regime(lower f) because of its lower stiﬀness. As a result, theoptimum cantilever depends on duration of the experiments, and the crossover between the two regimes
happens at surprisingly short times (∼25 ms).21
Our micromachined cantilever combines the advantages of the two diﬀerent cantilevers and improves
upon them. By decreasing the stiﬀness of a BioLever
Mini, the modiﬁed cantilever has a low-frequency
performance equal to an uncoated long BioLever. Thistranslates into excellent long-term force stability. In thethermally limited regime at higher f, the modiﬁed
cantilever has a lower PSD than the BioLever MiniD

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Figure 3. Comparing the performance of three diﬀerent
gold-coated cantilevers. Allan variance, a measure of precision over a given averaging time, was calculated from a
100 s trace measured 50 nm over a surface for a gold-coated
BioLever (BL) Mini (k = 79 pN/nm, gold), a gold coated long
BioLever (k = 5.0 pN/nm, purple), and a FIB-modiﬁed BL Mini(k = 5.1 pN/nm, lt. green). The gray dashed line is a referencewith a slope consistent with averaging Brownian motion.
The micromachined cantilever signiﬁcantly outperformed
both commercial cantilevers, particularly on time scales
longer than 0.1 s. On these time scales, the detrimental
eﬀects of gold on force stability are particularly pronounced. We note that at the very shortest times whenthe motion of the cantilever is correlated, the Allan varianceyields misleading results on force precision, and this regionof the trace is de-emphasized using a dotted gray line.

on longer time scales (0.150 s) because of the detrimental eﬀect of gold on force stability.24 Note that
performance of a gold-coated BioLever Mini deviatesfrom thermally limited performance at unexpectedly
short time scales (∼1 ms). Hence, in applications
requiring high reﬂectivity, our modiﬁed BioLever Mini
shows superior performance on both short and longtime scales.
Reduced Hydrodynamic Damping. For many biophysicists familiar with low Reynolds number hydrodynamics, a substantial reduction in β arising from achange in the shape, but not the longest length, ofan object is not obvious.41 On the other hand, experts
in cantilever dynamics may be familiar with priorsuccess in reducing β by FIB modification of a cantilever's shape.28,29 To quantify β, our first metric is basedon a classic result for an overdamped oscillator:41
β = k  τ, where τ is the characteristic relaxation time.
Our experimental definition of τ is when the autocorrelation dropped to 37% (e1) of its starting value. We
measured the autocorrelation for each cantilever nearand far from the surface (Figure 4). The resulting
autocorrelations were not a simple monotonic function but show a slight shoulder and, for some cantilevers, ringing. The shoulder is consistent with the firstharmonic of the cantilever. The ringing, or slightly
negative values at longer delay times, is consistent
with Q > 1. We note that the increased β near a surface
suppressed such ringing for both the long BioLeverand the modified BioLever Mini. Suppression of ringing
is advantageous in most SMFS applications, since a
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Figure 4. Temporal response of cantilevers in liquid near
(50 nm) and far (∼50 μm) from the surface. The autocorrelation of thermal motion as a function of time is plotted for
cantilevers near (colored) and far (black) from the surface foran uncoated long BioLever (top panel), an uncoated Biolever Mini (middle panel), and a modiﬁed BioLever Mini(bottom panel). A characteristic time for each cantilever isdetermined by the e1 point in the autocorrelation. A negative autocorrelation indicates the cantilever was slightly
underdampled (Q > 1). The data show all three cantilevers
with a longer correlation time and reduced Q near surfaces,
consistent with increased hydrodynamic drag due to theproximity of a surface. However, the modiﬁed BioLever Mini
shows less increase (85%) in its characteristic response
times in comparison to the other two cantilevers (250%
and 300% for the long Biolever and BioLever Mini, respectively). The characteristic response times far and near fromthe surface are 105 and 370 μs for the long BioLever, 8 and31 μs for the BioLever Mini and 20 and 37 μs for the modiﬁed
BioLever Mini. The stiﬀness for the three cantilevers are 6.4,
69, and 5.3 pN/nm for the long BioLever, the BioLever Mini,and the modiﬁed BioLever Mini, respectively.

sinusoidal variation of F complicates interpretation of athermally activated process, such as protein unfolding.
We ﬁrst analyzed the change in β when the cantilever is far from a surface on the basis of the ratioof β between an unmodiﬁed BioLever Mini and FIBmodiﬁed cantilever. This analysis yields a 6-fold reduction in β, a signiﬁcant change considering the length ofcantilever was not altered. We caution that this is aqualitative rather than a quantitative result. The calculation of β assumes overdamped motion, a valid
assumption for the modiﬁed, but not the unmodiﬁed,
BioLever Mini when far from the surface. Nonetheless,this result suggests a substantial reduction in β for themodiﬁed cantilevers.
The more relevant result is β when the cantilever is
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Figure 5. Improved single molecule force spectroscopy with FIB-modiﬁed cantilevers. (a) Cartoon of assay where eight
domains of a polyprotein are mechanically stretched by an AFM. (b) Forceextension records of mechanically unfolding apolyprotein of NuG2 at 400 nm/s with the long BioLever (red), the BioLever Mini (dark green), and the micromachined BioLever
Mini (lt. green). High-bandwidth data (50 kHz, gray) and smoothed data (1 kHz, colored) are shown. A worm-like-chain (WLC)
model (black line) well describes the stretching of the protein in one state (see also Figure S2a, SI). (c) Plot of residual force
ﬂuctuations after ﬁtting a 20 nm section of the 1 kHz data to a WLC model. The standard deviations of these ﬂuctuations are
4.1, 2.6, and 1.2 pN, using the same color scheme as in (b). Traces displaced vertically for clarity.

shows a 250 and 300% increase in τ and, therefore, β,respectively. In contrast, the increase in τ for the FIBmodiﬁed cantilever was only 85%. Hence, the large
central cutout of the cantilever not only reduced β far
from the surface, it also signiﬁcantly reduced the eﬀect
of increased β near surfaces due to the squeezed ﬁlm
damping, similar to prior results.28 To compare β for a
modiﬁed and a standard BioLever Mini at 50 nm overthe surface, we computed the ratio of β for the twocantilevers. This analysis shows a 12-fold reduction in β
for the individual cantilever shown in Figure 4. Thisratio is computed from records of the same individual
cantilever before and after modiﬁcation. The resulting
average reduction in β for multiple cantilevers was
10 ( 1 (N = 5). We note that both cantilevers showed anabsence of signiﬁcant ringing in their autocorrelation
function close to a surface (Figure 4). Hence, we expectthat this ratio in β to be a quantitative result.
For completeness, we also measured β using ageneralization of Stokes drag: F = β  v.41 To do so,
we started with the cantilever touching the surface andmoved it to 100 nm above the surface at increasing v
while measuring cantilever deﬂection. A plot of Fv forall three types of cantilevers studied showed a linear
relationship (Figure S1, Suppporting Information (SI)).
The slope of this line yields an independent estimate ofβ. The ratio of the slopes for the FIB-modiﬁed and astandard BioLever Mini is ∼5. However, we note that
this is not our preferred quantitative determination ofβ because of the dependence of β with height over thesurface and the short measurement time [1.6 ms(= 100 nm/60 μm/s)]. Summarizing two diﬀerent analyses show a signiﬁcant reduction in β near a surface,
the relevant regime for most AFM-based assays. Future
work can build upon our analysis for a more detailed
examination of β near a surface.
Improved Single Molecule Force Spectroscopy. To demonstrate the benefit of micromachined cantilevers for
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real-world applications, we performed a common
AFM-based single-molecule assay, mechanical unfolding of an individual protein (Figure 5a). The studied
protein consisted of 8 identical repeats of NuG2, acomputationally derived fast-folding variant of theprotein G B1 domain (GB1).42 As is typical for thesestudies, the cantilever was retracted at a constant v(400 nm/s) while measuring the resulting F. We recorded the data at 50 kHz and analyzed records thatshowed eight ruptures. The resulting traces showed
the classic sawtooth pattern for all three cantilevertypes studied (Figure 5b).
Visual inspection of Figure 5b shows a dramatic
improvement in the quality of the data by using a FIBmodiﬁed cantilever over either of the two commercial
cantilevers, particularly when looking at the records
smoothed to 1 kHz. We illustrate this improvement byanalyzing the ﬂuctuations in F. To do so, we ﬁt a short(20 nm) section of the smoothed forceextension
record to a worm-like chain (WLC) model and plotted
the residual ﬂuctuations in F (Figure 5c). The RMS
ﬂuctuations around mean position were 1.2 pN for
the FIB-modiﬁed BioLever while the unmodiﬁed BioLever mini and long BioLever showed 2.6 and 4.1 pN,
respectively. This reduction in force ﬂuctuations forthe modiﬁed BioLever Mini in a biophysical assay
conﬁrms a signiﬁcantly reduced β. More generally,
these forceextension curves were well ﬁt by a WLC
model (Figure S2a, SI) and illustrate the types ofimprovements aﬀorded by micromachined cantilevers
in a common AFM-based assay that requires shortterm force precision.
A 10-fold reduction in β improves SMFS in anothermanner; it decreases the hydrodynamic force applied
to the cantilever when collecting forceextension data
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Figure 6. Micromachined cantilever shows improved performance while stretching a protein. (a) Schematic of theexperiment showing a protein stretched at constant extension. (b) Force-versus-time records for three diﬀerent cantilevers
stretching a protein to F ≈ 50 pN while the stage position was held ﬁxed. High-bandwidth data (50 kHz, gray) and smoothed
data (1 kHz, colored) are shown. Traces displaced for clarity. The standard deviation for the 1 kHz data was 5.2, 3.2, and 1.0 pN,
for the long BioLever (red), BioLever Mini (dark green), and the FIB-modiﬁed BioLever Mini (lt. green), respectively. The
stiﬀnesses for the three cantilevers were 4.4, 74, and 5.9 pN/nm in the same order. (c) Schematic showing the abrupt unfoldingof a protein observed while holding the cantilever at constant height over the surface. (d) Averaged force-versus-time records
show the step response function of the diﬀerent cantilevers. The characteristic times for the three cantilevers were 450 ((40),53 ((16), and 76 ((18) μs for a long BioLever (red), a BioLever Mini (dark green), and a modiﬁed BioLever Mini (lt. green),
respectively. Note, the smoother appearance of the long BioLever record relative to the two other cantilevers is an artiﬁcial
result arising from plotting data at 50 kHz where the individual points are not statistically independent (see also Figure 3).

low as a few μm/s when using standard cantilevers.43,44
Reduction in hydrodynamic force also facilitates SMFS
at very high pulling velocities (v = 0.55 mm/s), which
has recently opened the door for direct comparison between SMFS data and molecular dynamics
simulations.45
Cantilevers with reduced β can improve imaging
applications as well, in particular nanoscale mapping of
the mechanical properties of materials.46 In such experiments, an array of forcedistance curves is measured at every xy point, a so-called forcevolume orforcedistance map. This process is usually very slow.
Recently, software and hardware enhancements have
led to rapid nanomechanical investigations of cells47
and membrane proteins with molecular48 and submolecular resolution.49 Rapid force-volume imaging
is commercially available [e.g., PeakForce Tapping(Bruker)]. The quality and the speed of such experiments could be improved by using cantilevers thathave both reduced k and β.
Enhanced Sub-pN Precision and Stability in Surface-AnchoredBiophysical Assay. Equilibrium studies of macromolecular folding need excellent short-term force precisioncoupled with long-term stability. To probe long-term
stability in a single-molecule assay, we first mechanically unfolded a polyprotein of NuG2 to its full extension and then lowered the cantilever until F = 50 pN.
This F prevented refolding while allowing for theacquisition of a 100 s trace recorded at a stationary
BULL ET AL.

stage position (Figure 6a). The resulting data showed
excellent force stability (Figure 6b) when the instrument had been allowed to settle for 12 h after
cantilever mounting. Quantitatively, the FIB-modified
cantilever showed an excellent combination of force
precision and force stability with an integrated force
noise of 1.1 pN over five decades of bandwidth (Δf =0.011000 Hz). The same metric yielded 2.4 and 5.2 pN
for the BioLever Mini and the long BioLever, respectively. We note that this level of performance relied
upon the stability of our commercial AFM; neither F nor
the tipsample separation25 was actively stabilized.
Rather, a closed-loop piezo-electric stage was used tomaintain the stage position, as is typical in such assays.
Different instruments will yield different results. However, the critical point is that micromachined cantilevers maintained essentially the same force stability
in a protein-based assay as for an isolated cantilever(Figure 1e). As a result, our ∼50-fold improvement inbandwidth for sub-pN force precision is preserved in abiophysical assay even when the cantilever is anchored
to a surface through the molecule under study.
These long records of protein extension also allowed us to explore short-term force precision in anassay analogous to equilibrium studies of protein
folding and unfolding.15 As discussed above, better
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Figure 7. Analyzing the limits for detection of short-lived events as a function of force and event duration (Δt).
(a, inset) Forceextension record for stretching a polyprotein consisting of three (orange) and four (blue) unfolded NuG2
domains. The two curves were considered distinguishable, and therefore color coded, when they were separated by twice thestandard deviation in the force noise in data smoothed over the speciﬁed event time. (a) Plot of applied force versus resolvablechange in contour length (ΔL) for a 1 ms event for the FIB-modiﬁed cantilever (green) and the uncoated long BioLever (red),
the two cantilevers with necessary force stability for long equilibrium experiments. (b) Same as in panel (a) but analyzed
assuming a 10 ms event. Note that the FIB-modiﬁed cantilever can resolve a smaller change in ΔL during a 1 ms event than thelong BioLever can in a 10 ms event.

for the isolated modiﬁed BioLever versus one pullingon a protein at constant extension shows similar performance (Figure S3, SI). This analysis shows that improvements in short-term force precision aﬀorded by
the micromachined cantilever in a biophysical assay
continued to nearly match that of an isolated cantilever.
Sensitive but Responsive in Protein-Unfolding Studies. Improved protein folding and unfolding studies rely not
only on force precision, but also on cantilevers that
respond rapidly to an abrupt change in applied F. While
short-term force noise is governed by β, the mechanical response time is given by τ = β/k in the overdamped limit. Thus, in general, softer cantilevers areless responsive at fixed β.
We used the abrupt opening of a protein to directly
measure the response function of the cantilever(Figure 6c). Our protocol was similar to the proteinpulling protocol used for Figure 5. In this revised assay,
we selected for a full-length construct, lowered theextension until F ≈ 10 pN and then kept the stage at aconstant position. After waiting 5 s, the extension was
rapidly increased until F ≈ 70 pN. The resulting data
was then recorded at high bandwidth (50 kHz) for the
next 3 s. If the tether did not break, the force wasreturned to 10 pN for 5 s and the process repeated. To
improve the signal-to-noise ratio, we averaged several unfolding trajectories (N = 6). The resulting data
showed an approximately exponential drop in F due tothe abrupt unfolding of a protein (Figure 6d). The
characteristic times for a normal BioLever Mini and a
modiﬁed BioLever Mini were 53 ((16) and 76 ((18) μs,
respectively. These times were signiﬁcantly faster than
the 450 ((40) μs measured for a long BioLever. Thus,
our soft cantilevers facilitate equilibrium studies because of their long-term force stability, while their
reduced β allows them to be responsive in detecting
short-lived events.
BULL ET AL.

Improved Detection of Short-Lived States. Short-term
force precision facilitates many nanoscience applications. For instance, better short-term force precisionenables more robust detection of transiently populated states during SMFS studies of protein folding.
Such transient states may last only 1 ms or less inequilibrium studies.37 Transiently populated unfolding
intermediates are also observed while mechanicallyunfolding proteins26,27 and RNA22 at a constant velocity. To analyze the limits in detecting the folding andunfolding of proteins in an equilibrium assay, we
calculated the smallest protein-unfolding event that
could be resolved when using the two cantilevers thatshow the necessary long-term force stability for suchan assay, the modified BioLever Mini and the uncoatedlong BioLever. A change in L was considered detectable if two states, defined by WLC curves, were separated by twice the standard deviation in the data ina given time interval where the initial L was 53 nm(Figure 7a, inset).
Detection of short-lived events depends on theapplied force and the duration of the event. We used
10 pN as a representative force, since equilibrium
folding between states for a variety of biomolecules
is seen at 512 pN.15,37,38,50 We initially used thestandard deviation of a cantilever's motion whensmoothed over a 1 ms period. This analysis showedthat our modiﬁed cantilever could detect a ΔL = 13
amino acids [aa (1 aa = 0.36 nm)] in 1 ms (Figure 7a). In
comparison, the same analysis yielded ΔL = 31 aa for
an uncoated long BioLever. Longer events allow for
more averaging of Brownian motion, and therefore asmaller ΔL is resolvable. When a 10 ms event was
analyzed, the micromachined cantilever and the long
BioLever could detect a ΔL of 6 and 18 aa, respectively(Figure 7b). We note this analysis is based solely on
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For comparison, we have also FIB-modiﬁed short BioLevers [L = 60 μm; k = 30 pN/nm (unmodiﬁed values)].
Unfortunately, these cantilevers rarely unfolded, making them a poor choice for routine use.
Our micromachined cantilevers showed long-term
stability equal to an uncoated long BioLever.24 However, this level of performance typically takes 12 h toattain, unlike uncoated cantilevers, which attained
sub-pN stability just 30 min after mounting.24 We
emphasize that this limitation will only aﬀect long timescale experiments. The improved short-term force
precision (Figure 5) is immediately accessible anduseful in a wide range of AFM-based assays. For
scientists familiar with standard gold-coated cantilevers, the long-term drift of our modiﬁed cantilevers
is signiﬁcantly less and stabilizes on a more rapid timescale than traditional cantilevers.

METHODS

The protocol begins by loading the cantilevers into the FIB
using a custom-made metal mounting plate. This mounting
geometry held 9 cantilevers, which were aﬃxed to the mounting plate with graphite tape. The graphite tape helped minimize
the charging of the cantilever during FIB-modiﬁcation; excess
charge on insulating samples leads to degraded performancein scanning electron microscopy (SEM) and FIB applications.51
We next loaded the sample into the FIB. It took ∼20 min to

Focused Ion Beam Modification of a Commercial Cantilever. We used
a dual-beam FIB (Nova NanoLab 600, FEI) that has both anelectron (e) beam and an ion (Ga2þ) beam. The e-beam
was used to write the glass-like structure, to compensate forcharging during FIB-milling,51 and to image the resulting
structure. The ion-beam was used to remove material fromthe cantilever.

BULL ET AL.

ARTICLEthe comparatively slow mechanical relaxation of thelong BioLever (0.45 ms), which makes a 1 ms event
even more challenging to resolve.
Dynamic experiments also show folding intermediates26,27 and beneﬁt from improved detectionof short-lived events. Unfolding forces of >50 pN at
pulling rates of 300500 nm/s are common.26,27 In thisregime of ∼23 ms/nm at F = 50 pN, our analysis
suggests that detecting even a single amino acidchange (ΔL = 1 aa) is possible with FIB-modiﬁed
cantilevers.
In summary, our modiﬁed cantilevers excel at resolving small changes in ΔL over short periods. Indeed,
they can resolve a smaller ΔL in 1 ms than an uncoatedlong BioLever can over 10 ms without sacriﬁcing
the force stability of the uncoated cantilever. Moreover, these modiﬁed cantilevers respond more quickly
and have a high reﬂectivity. Overall, we expect these
cantilevers will aid both equilibrium and dynamic
studies of protein folding by AFM.
Cost, Usability, and Limitations. Our FIB-modified cantilevers are not too costly for routine use. After an
extended research and development phase, our marginal cost for making these cantilevers in batches of 9 is∼$30/ea at a typical academic rate for using an FIB($75/h) operated by an undergraduate research assistant ($12/h). This marginal cost is about two-thirds thecost of a BioLever Mini or one-third of the cost of a
BioLever Fast. Thus, for a relatively modest increase incost per cantilever, one gets a dramatic improvementin performance over a broad bandwidth.
Perhaps more important than the marginal cost isthat we reused FIB-modiﬁed cantilevers in SMFS applications over multiple days (Figure S2b, SI). Unlike
imaging applications, SMFS does not require a particularly sharp tip. After cleaning, the reused cantilevershowed no change in stiﬀness or loss in performance,increasing the return on investment in fabricating them.
Like many soft cantilevers, these modiﬁed cantilevers sometimes folded when immersed into liquid.
However, given suﬃcient time (∼5 min), they typicallyunfolded. If the cantilever remained folded, dewetting/rewetting a few times typically induced unfolding. In
cases where rewetting was unsuccessful, the cantilevers were induced to unfold by brieﬂy dipping themin ethyl alcohol. In our experience, this behavior
is quite comparable to standard long BioLevers.

CONCLUSIONS
We have developed an eﬃcient way to fabricate soft,
but short, AFM cantilevers. Their 10-fold lower β leads
to better short-term force precision. Their 10-fold lowerk, coupled with the removal of all of the gold except
from the end of the cantilever, leads to excellent longterm force stability. The protected gold patch enableshigh reﬂectivity, without loss of such stability. Importantly, these FIB-modiﬁed cantilevers are neither too
complex to fabricate nor too costly for routine use in acommercial AFM. Moreover, our work extends earlier
thermal noise characterization of FIB-modiﬁed cantilevers28,29 by directly demonstrating enhanced performance in a common single-molecule assay, the mechanical unfolding of a protein. We highlighted thisimprovement in three diﬀerent force spectroscopy
assays. First, we showed signiﬁcantly improved shortterm force precision during a dynamic unfolding assay(Figure 5). Second, we showed sub-pN performanceover ﬁve decades of bandwidth (Δf ≈ 0.011000 Hz)
while stretching a surface-anchored protein (Figure 6b).
Finally, we showed the temporal response of the cantilever to a protein unfolding remained excellent (∼70 μs)
despite the cantilever's reduced k (Figure 6d). Hence,these FIB-modiﬁed cantilevers show enhanced performance in multiple force spectroscopy assays. We expect
these enhancements can immediately improve a wide
range of other AFM-based studies in biophysics andnanotechnology.

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as evidenced by the lighter color gray regions shown in
Figure 1c. We observed no noticeable detrimental eﬀects
of charging when moving the ion beam in a cyclic patternrepeated every ∼0.5 s. We repeated this pattern a total of
500 times over 4 min.
We removed the cantilever's gold coating using a ∼40 s wet
etch (Type TFA, Transene) (Figure 2e). The underlying chromium
adhesion layer was then removed using a ∼40 s wet etch
(Cr Etchent, Transene). Each etch was separated by a water
bath and blotted dry using the corner of a kimwipe (Kimtech).
Characterizations of Micromachined Cantilevers. We used a commercial AFM (Cypher, Asylum Research with its standard, notsmall, spot size module installed) to compare the performanceof the modified cantilevers to commercial ones that had theirgold coating removed. For these measurements, the AFM wasconfigured using the instrument's “crosspoint panel” to bypass
the default high-pass filter, enabling accurate measurement
over a broad frequency range. Cantilever stiffness was calibrated in liquid far from the surface using the instrument's builtin fitting algorithm.55 To determine the reduction in k due to FIB
modification, the stiffness of all micromachined cantilevers wasmeasured before and after modification.
We used two primary metrics to assess cantilever performance in liquid: the integrated force noise and the Allan
variance.39 After letting each cantilever equilibrate in 150 mM
phosphate buﬀer (pH 8) for 12 h, we touched the cantilever oﬀ
the surface and then retracted it by 50 nm. The cantilever's
thermal motion was then digitized in ﬁve 100 s segments at
50 kHz. We ﬁrst calculated the positional power spectral density(PSD) and then multiplied it by the cantilever stiﬀness to yield
the force PSD. The integrated force noise was calculated byintegrating the force power spectral density (PSD) over the
speciﬁed bandwidth. The Allan variance σ was calculated fromthe same data using
rﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
1
σx (T) ¼
Æ(xi þ 1  xi )2 æT(1)
2

ARTICLE

achieve a vacuum of 105 Torr (1.3 mPa), our typical basepressure.
We used an e-beam induced deposition process52 to form
the capping layer (Figure 2b) that protected the gold during
subsequent chemical etches. The mask was transparent andmade from a precursor gas (TEOS, FEI). The gas was introduced
into the FIB vacuum chamber through a micromanipulatorcontrolled needle as part of the manufacturer's gas injection
system (GIS). This system allowed us to position a needle close(∼400 μm) to the sample, improving gas adsorption withoutcompromising the base pressure of the main FIB chamber. The
gas ﬂow rate was controlled by the pressure of the precursor
gas, which was typically ∼1 Torr (130 Pa). Gas adsorbed on thesurface forms a solid glass-like material when irradiated with ane-beam of adequate energy (>1 eV). The precursor gas is more
eﬀectively activated by secondary electrons (∼10 eV) than highenergy electrons (10 keV), due to its larger cross section for low
energy electrons. Secondary electrons (<50 eV) are eﬃciently
produced by the e-beam as it scatters in the substrate (i.e., thesilicon nitride of the cantilever).52 To do so, we used a tightly
focused (∼150 nm diameter) 100-pA 5 keV e-beam. Rasterscanning this e-beam enabled us to generate the desired
8  12 μm2 pattern. The pattern itself was laid out using the
manufacturer's software and had a pixel size of the ∼175 nm.
The dwell time per pixel during the writing process was 0.04 ms.
We repeated the pattern 5000 times for a total write time of 6 min.
We used the e-beam limited regime for deposition, in contrast to
an adsorption-limited one, to achieve better mask continuity andimproved deposition purity.52
To remove the large, central region of the cantilever(Figure 2c), we ﬁrst had to overcome a signiﬁcant problem
associated with FIB modiﬁcation of soft cantilevers: they vibrated and/or folded back upon themselves during the milling
process. The cause of these unwanted cantilever dynamics was
charging of the cantilever, a problem that was partially, but not
fully, alleviated by using graphite tape in the mounting process.
The traditional solution to this problem is to sputter extra gold
onto the cantilever,51 an added step in handling of eachcantilever. We avoided this additional step with two modiﬁcations to the FIB-milling process. The ﬁrst modiﬁcation was toapply a defocused e-beam (10 μm diam; 1.5 nA) to the cantileverconcurrent with the FIB milling. The negative electrons helped
neutralize the positive charge build up from the Ga2þ ions.51
The second modiﬁcation was to start the milling process on thesegment of the cantilever farthest from the chip. This simple
procedure preserved the inherent stiﬀness of the unmodiﬁed
cantilever for as long as possible to resist folding or vibrating.
The actual milling was accomplished using a relatively tightly
focused ion beam (<300 nm diam; 100 pA) to cut the perimeteraround a rectangular region. To avoid damage to the cantilever
during the prolonged irradiation resulting from the focusing
process,53 we focused the ion beam on a triangular portion of
the cantilever material left over from the manufacturing processthat is next to the rectangular cantilever as seen in Figure 1b.
After focusing, the cantilever thickness was milled throughentirely in ∼5 min using a 40 nm pixel spacing and a ∼0.2 ms
dwell times. At the completion of cutting, the rectangularsection fell away, leaving two narrow supports holding the
tip-containing section of the cantilever (Figure 2c).
To further reduce k, the narrow-width supports werethinned by ∼30%. The primary challenge was maintaining
cantilever integrity. We achieved better results when we implemented a technique using an ion beam defocused to 2 μm,
a value chosen on the basis of Stokes et al.51 This width wasslightly wider than the full width of the support. Two potential
explanations for this improved performance are (i) a larger
beam size should reduce localized charging that could distortthese fragile cantilevers53 and (ii) a reduced dosage per unit
area should reduce the rate of milling, making the thinningmore uniform over the width of the support.54 Once we deﬁned
this large beam size, the 100 pA beam was moved in a linear
pattern along the support in a single-pixel-wide line pattern,removing the exposed top layer of material. To minimize excess
milling at the turning points in the line pattern, the turning
points were chosen to be outside the narrow support region,

where T is the averaging time interval, and xi is the mean valueof the data over the ith time interval.39 We have previously used
these metrics to analyze uncoated BioLever Minis and long
BioLevers.21,24
We used two methods to probe the hydrodynamic dragof the cantilever. The ﬁrst was quite simple, but assumed
overdamped motion. An overdamped oscillator displaced
from equilibrium will decay exponentially toward equilibrium.
The characteristic decay time is given by τ = β/k.41 We haveexperimental access to τ and k, allowing β to be computed.
The stiﬀness calibration yielded k. We computed τ basedon e1 value in the autocorrelation in the cantilever's thermal
motion.
The second method is based upon Stokes drag, where thecantilever was moved vertically up and down in a sawtoothpattern at varying velocities (v = 1560 μm/s). A uniform
velocity led to a constant cantilever deﬂection. A generalizationof Stoke's law yields a phenomenological deﬁnition of hydrodynamic drag, β = F/v. From these measurements, we got asecond determination of β for the three diﬀerent cantilevers
studied (Figure S1, SI).
Single Molecule Force Spectroscopy of a Polyprotein. To test our
micromachined cantilevers in a biological assay, we studied a
polyprotein of NuG2, a fast-folding variant of GB1.42 In thisconstruct, there are eight identical tandem repeats of the NuG2
domain, a N-terminal His-tag to facilitate purification, and an
N-terminal cysteine to facilitate surface anchoring.56 The mechanical unfolding properties of this polyprotein have beenpreviously characterized using AFM-based SMFS.56
Historically, AFM-based SMFS is done by passively absorbing the protein onto a glass or mica surface and relying on
nonspeciﬁc interactions between the tip and the protein. To
improve the stability of the attachment for taking 100 s records,
we prepared glass substrates coated with NHS-PEG-maleimide
based upon prior work.57 The PEG surface dramatically decreases the nonspeciﬁc sticking of the protein to the surface.58
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Acknowledgment. We thank G. King for useful discussions,A. Adhikari for help in preparing functionalized glass surfaces,
and B. Baxley for scientiﬁc illustration. This work was supported by
the National Science Foundation [DBI-0923544, Phys-1125844],
and NIST. Mention of commercial products is for information only;
it does not imply NIST's recommendation or endorsement. T.T.P. isa staﬀ member of NIST's Quantum Physics Division.
Supporting Information Available: Figures showing hydrodynamic drag of cantilevers near a surface, forceextension
data modeled with WLC ﬁts, forceextension data taken withreused FIB-modiﬁed cantilevers, and comparison of Allan variance when pulling on a protein unfolding with that of an
isolated cantilever. This material is available free of charge via
the Internet at http://pubs.acs.org.

BULL ET AL.

REFERENCES AND NOTES
1. Binnig, G.; Quate, C. F.; Gerber, C. Atomic Force Microscope. Phys. Rev. Lett. 1986, 56, 930–933.
2. Engel, A.; Gaub, H. E. Structure and Mechanics of Membrane Proteins. Annu. Rev. Biochem. 2008, 77, 127–148.
3. Rief, M.; Gautel, M.; Gaub, H. E. Unfolding Forces of Titin
and Fibronectin Domains Directly Measured by AFM. Adv.
Exp. Med. Biol. 2000, 481, 129–141.
4. Muller, D. J.; Helenius, J.; Alsteens, D.; Dufrene, Y. F. Force
Probing Surfaces of Living Cells to Molecular Resolution.
Nat. Chem. Biol. 2009, 5, 383–390.
5. Muller, D. J.; Dufrene, Y. F. Atomic Force Microscopy as a
Multifunctional Molecular Toolbox in Nanobiotechnology.
Nat. Nanotechnol. 2008, 3, 261–269.
6. Muller, D. J.; Sapra, K. T.; Scheuring, S.; Kedrov, A.; Frederix,
P. L.; Fotiadis, D.; Engel, A. Single-Molecule Studies ofMembrane Proteins. Curr. Opin. Struct. Biol. 2006, 16,
489–495.
7. Scheuring, S.; Sturgis, J. N. Chromatic Adaptation of Photosynthetic Membranes. Science 2005, 309, 484–487.
8. Ando, T. High-Speed Atomic Force Microscopy Coming ofAge. Nanotechnology 2012, 23, 062001.
9. Fernandez, J. M.; Li, H. Force-Clamp Spectroscopy Monitors the Folding Trajectory of a Single Protein. Science
2004, 303, 1674–1678.
10. Rief, M.; Gautel, M.; Oesterhelt, F.; Fernandez, J. M.; Gaub,
H. E. Reversible Unfolding of Individual Titin Immunoglobulin Domains by AFM. Science 1997, 276, 1109–1112.
11. Muller, D. J.; Schabert, F. A.; Buldt, G.; Engel, A. Imaging Purple Membranes in Aqueous Solutions at SubNanometer Resolution by Atomic Force Microscopy. Biophys. J. 1995, 68, 1681–1686.
12. Schabert, F. A.; Henn, C.; Engel, A. Native Escherichia coli
OmpF Porin Surfaces Probed by Atomic Force Microscopy.
Science 1995, 268, 92–94.
13. Muller, D. J.; Fotiadis, D.; Scheuring, S.; Muller, S. A.; Engel,
A. Electrostatically Balanced Subnanometer Imaging ofBiological Specimens by Atomic Force Microscope. Biophys. J. 1999, 76, 1101–1111.
14. Muller, D. J.; Schoenenberger, C. A.; Schabert, F.; Engel, A.
Structural Changes in Native Membrane Proteins Monitored at Subnanometer Resolution with the Atomic ForceMicroscope: A Review. J. Struct. Biol. 1997, 119, 149–157.
15. Junker, J. P.; Ziegler, F.; Rief, M. Ligand-Dependent Equilibrium Fluctuations of Single Calmodulin Molecules.
Science 2009, 323, 633–637.
16. Rief, M.; Clausen-Schaumann, H.; Gaub, H. E. SequenceDependent Mechanics of Single DNA Molecules. Nat.
Struct. Biol. 1999, 6, 346–349.
17. Rief, M.; Oesterhelt, F.; Heymann, B.; Gaub, H. E. Single
Molecule Force Spectroscopy on Polysaccharides by
Atomic Force Microscopy. Science 1997, 275, 1295–1297.
18. Muller, D. J.; Dufrene, Y. F. Atomic Force Microscopy: A
Nanoscopic Window on the Cell Surface. Trends Cell Biol.
2011, 21, 461–469.
19. Dufrene, Y. F.; Evans, E.; Engel, A.; Helenius, J.; Gaub, H. E.;
Muller, D. J. Five Challenges to Bringing Single-Molecule
Force Spectroscopy into Living Cells. Nat. Methods 2011, 8,
123–127.
20. Casuso, I.; Scheuring, S. Automated Setpoint Adjustment
for Biological Contact Mode Atomic Force Microscopy
Imaging. Nanotechnology 2010, 21, 035104.
21. Sullan, R. M.; Churnside, A. B.; Nguyen, D. M.; Bull, M. S.;
Perkins, T. T. Atomic Force Microscopy with Sub-PicoNewton
Force Stability for Biological Applications. Methods 2013, 60,
131–141.
22. Liphardt, J.; Onoa, B.; Smith, S. B.; Tinoco, I. J.; Bustamante,
C. Reversible Unfolding of Single RNA Molecules by Mechanical Force. Science 2001, 292, 733–737.
23. Viani, M. B.; Schaﬀer, T. E.; Chand, A.; Rief, M.; Gaub, H. E.;
Hansma, P. K. Small Cantilevers for Force Spectroscopy ofSingle Molecules. J. Appl. Phys. 1999, 86, 2258–2262.
24. Churnside, A. B.; Sullan, R. M.; Nguyen, D. M.; Case, S. O.;
Bull, M. S.; King, G. M.; Perkins, T. T. Routine and Timely SubpicoNewton Force Stability and Precision for BiologicalVOL. XXX’

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NuG2 polyprotein to the surface via its N-terminal cysteine. We
incubated 1 μL of ∼500 nM NuG2 in 100 μL of 150 mM
phosphate buﬀer (pH 8) and 1 μL of 10 mM TCEP for 1 h atroom temperature and then rinsed with 400 μL of phosphatebuﬀer. Cantilevers were illuminated with UV light for 1 min(Bondwand, Electro-Lite Corp.) immediately prior to loading topromote nonspeciﬁc attachment to the uncoated cantilever.
Samples and the cantilever were loaded and equilibrated for >1
h prior to measurements. All experiments were done in 150 mM
phosphate buﬀer with the AFM's temperature stabilized
to 27 C.
Data were recorded with a custom program written in
Asylum's MacroBuilder software. We initiated nonspeciﬁc attachment of the protein to the tip by pressing the tip into thesample at 200 pN for 1 s. The stage was then retracted at v =400 nm/s. The records were digitized at 50 kHz. About 1% of theattempts showed a sawtooth pattern consisting of 8 ruptureswith a change in contour length consistent with the unfoldingof a NuG2 domain (ΔL ≈ 17.7 nm) (Figure S2a, SI).59 These
records were selected for analysis. For display, these highbandwidth records were boxcar-averaged and decimated to a
1 kHz data rate. To quantify the force noise during a pullingrecord, a short (20 nm) linear section of the 1 kHz forceextension record was ﬁt to a WLC model. The residuals from this ﬁtwere then plotted (Figure 5c).
To test for long-term force stability, we used a real-time
selection process for molecules showing unfolding of all eight
domains and a strong tip-protein interaction. This selection was achieved by proceeding only with molecules thatshowed F > 150 pN at an extension of at least 130 nm andthen rapidly reversing the direction of the stage until F = 50 pN.
The stage was held stationary during subsequent 100 s
while the cantilever deﬂection was recorded at 50 kHz. At
the conclusion of the 100 s recording, we reconﬁrmed thepresence of the biomolecule by pulling the molecule until
rupture. These trials were completed on a micromachined
BioLever Mini as well as on an uncoated long BioLever and a
BioLever Mini.
The temporal resolution of each cantilever was monitored
by looking at the force decay after the unfolding of a NuG2
domain. As above, we selected for molecules exhibiting F > 150 pN
and an extension greater than 130 nm. In this assay, the stage
was rapidly moved backward a precalculated ﬁxed amount(40 nm), suﬃcient to decrease F to ∼10 pN based on a WLC model.
After waiting for 5 s, we abruptly moved the stage position until
F = 70 pN. We measured the subsequent cantilever motion for 3 s
at 50 kHz. If the attachment to the polyprotein did not rupture,then the process was repeated multiple times. Individual portionsof records consistent with a NuG2 domain were aligned andaveraged. An exponential ﬁt to this average curve yielded thetime constant of the mechanical relaxation.
A micromachined cantilever could be reused in SMFS assays
over multiple days. After each use, the cantilever was rinsed inultrapure water and then blotted dry. At a later time, thecantilever was plasma cleaned (25 SCCM O2, 10 s, 250 W) using
a standard plasma cleaner (PlasmaSTAR, Axic). Cantilevers werestored in a gel pack for later reuse.
Conflict of Interest: The authors declare no competing
ﬁnancial interest.

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26.

27.

28.

29.
30.
31.

32.
33.
34.
35.
36.

37.

38.
39.
40.
41.
42.
43.
44.

45.

46.

BULL ET AL.

47.

48.

49.
50.
51.

52.
53.
54.

55.

56.
57.

58.

59.

by Force-Distance Curve-Based AFM. Nat. Methods 2013,10, 847–854.
Alsteens, D.; Dupres, V.; Yunus, S.; Latge, J. P.; Heinisch, J. J.;
Dufrene, Y. F. High-Resolution Imaging of Chemical andBiological Sites on Living Cells using Peak Force Tapping
Atomic Force Microscopy. Langmuir 2012, 28, 16738–
16744.
Medalsy, I.; Hensen, U.; Muller, D. J. Imaging and Quantifying Chemical and Physical Properties of Native Proteins at
Molecular Resolution by Force-Volume AFM. Angew.
Chem., Int. Ed. 2011, 50, 12103–12108.
Rico, F.; Su, C. M.; Scheuring, S. Mechanical Mapping ofSingle Membrane Proteins at Submolecular Resolution.
Nano Lett. 2011, 11, 3983–3986.
Cecconi, C.; Shank, E. A.; Bustamante, C.; Marqusee, S.
Direct Observation of the Three-State Folding of a Single
Protein Molecule. Science 2005, 309, 2057–2060.
Stokes, D. J.; Vystavel, T.; Morrissey, F. Focused Ion Beam
(FIB) milling of Electrically Insulating Specimens Using
Simultaneous Primary Electron and Ion Beam Irradiation.
J. Phys. D: Appl. Phys. 2007, 40, 874–877.
van Dorp, W. F.; Hagen, C. W. A Critical Literature Review of
Focused Electron Beam Induced Deposition. J. Appl. Phys.
2008, 104, 081301.
Giannuzzi, L. A.; Stevie, F. A. Introduction to Focused Ion
Beams: Instrumentation, Theory, Techniques and Practice;Springer: New York, 2005.
Hopman, W. C. L.; Ay, F.; Hu, W. B.; Gadgil, V. J.; Kuipers, L.;
Pollnau, M.; de Ridder, R. M. Focused Ion Beam Scan
Routine, Dwell Time and Dose Optimizations for Submicrometre Period Planar Photonic Crystal Components
and Stamps in Silicon. Nanotechnology 2007, 18, 195305.
Proksch, R.; Schaﬀer, T. E.; Cleveland, J. P.; Callahan, R. C.;
Viani, M. B. Finite Optical Spot Size and Position Corrections in Thermal Spring Constant Calibration. Nanotechnology 2004, 15, 1344–1350.
Cao, Y.; Kuske, R.; Li, H. Direct Observation of Markovian
Behavior of the Mechanical Unfolding of Individual Proteins. Biophys. J. 2008, 95, 782–788.
Zimmermann, J. L.; Nicolaus, T.; Neuert, G.; Blank, K. ThiolBased, Site-Speciﬁc and Covalent Immobilization of Biomolecules for Single-Molecule Experiments. Nat. Protoc.
2010, 5, 975–985.
Ha, T.; Rasnik, I.; Cheng, W.; Babcock, H. P.; Gauss, G. H.;
Lohman, T. M.; Chu, S. Initiation and Re-Initiation of DNA
Unwinding by the Escherichia coli Rep Helicase. Nature
2002, 419, 638–641.
Cao, Y.; Balamurali, M. M.; Sharma, D.; Li, H. B. A Functional
Single-Molecule Binding Assay via Force Spectroscopy.
Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 15677–15681.

VOL. XXX’

NO. XX’

000–000’

ARTICLE

25.

Applications of Atomic Force Microscopy. Nano Lett. 2012,12, 3557–3561.
King, G. M.; Carter, A. R.; Churnside, A. B.; Eberle, L. S.;
Perkins, T. T. Ultrastable Atomic Force Microscopy: AtomicScale Lateral Stability and Registration in Ambient Conditions. Nano Lett. 2009, 9, 1451–1456.
Zocher, M.; Roos, C.; Wegmann, S.; Bosshart, P. D.; Dotsch,
V.; Bernhard, F.; Muller, D. J. Single-Molecule Force Spectroscopy from Nanodiscs: An Assay to Quantify Folding,
Stability, and Interactions of Native Membrane Proteins.
ACS Nano 2012, 6, 961–971.
Sapra, K. T.; Besir, H.; Oesterhelt, D.; Muller, D. J. Characterizing Molecular Interactions in Diﬀerent Bacteriorhodopsin Assemblies by Single-Molecule Force Spectroscopy.
J. Mol. Biol. 2006, 355, 640–650.
Maali, A.; Cohen-Bouhacina, T.; Jai, C.; Hurth, C.; Boisgard,R.; Aime, J. P.; Mariolle, D.; Bertin, F. Reduction of the
Cantilever Hydrodynamic Damping Near a Surface by
Ion-Beam Milling. J. Appl. Phys. 2006, 99, 024906.
Hodges, A. R.; Bussmann, K. M.; Hoh, J. H. Improved Atomic
Force Microscope Cantilever Performance by Ion Beam
Modiﬁcation. Rev. Sci. Instrum. 2001, 72, 3880–3883.
Noy, A.; Vezenov, D. V.; Kayyem, J. F.; Meade, T. J.; Lieber,
C. M. Stretching and Breaking Duplex DNA by Chemical
Force Microscopy. Chem. Biol. 1997, 4, 519–527.
Denis, F. A.; Hanarp, P.; Sutherland, D. S.; Gold, J.; Mustin, C.;
Rouxhet, P. G.; Dufrene, Y. F. Protein Adsorption on Model
Surfaces with Controlled Nanotopography and Chemistry.
Langmuir 2002, 18, 819–828.
Piner, R. D.; Zhu, J.; Xu, F.; Hong, S.; Mirkin, C. A. “Dip-Pen”
Nanolithography. Science 1999, 283, 661–663.
Liu, M. Z.; Amro, N. A.; Chow, C. S.; Liu, G. Y. Production of
Nanostructures of DNA on Surfaces. Nano Lett. 2002, 2,
863–867.
Labuda, A.; Bates, J. R.; Grütter, P. H. The Noise of CoatedCantilevers. Nanotechnology 2012, 23, 025503.
Svoboda, K.; Block, S. M. Biological Applications of Optical
Forces. Annu. Rev. Biophys. Biomol. Struct. 1994, 23, 247–
285.
Kawakami, M.; Taniguchi, Y.; Hiratsuka, Y.; Shimoike, M.;
Smith, D. A. Reduction of the Damping on an AFM
Cantilever in Fluid by the Use of Micropillars. Langmuir
2010, 26, 1002–1007.
Yu, H.; Liu, X.; Neupane, K.; Gupta, A. N.; Brigley, A. M.;
Solanki, A.; Sosova, I.; Woodside, M. T. Direct Observationof Multiple Misfolding Pathways in a Single Prion Protein
Molecule. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 5283–
5288.
Stigler, J.; Ziegler, F.; Gieseke, A.; Gebhardt, J. C. M.; Rief, M.
The Complex Folding Network of Single Calmodulin Molecules. Science 2011, 334, 512–516.
Sullivan, D. B.; Allan, D. W.; Howe, D. A.; Walls, E. L.
Characterization of Clocks and Oscillators; U.S. Government Printing Oﬃce: Washington, D.C., 1990.
Czerwinski, F.; Richardson, A. C.; Oddershede, L. B. Quantifying Noise in Optical Tweezers by Allan Variance. Opt.
Express 2009, 17, 13255–13269.
Howard, J. Mechanics of Motor Proteins and the Cytoskeleton; Sinauer Associates: Sunderland, MA, 2001.
Nauli, S.; Kuhlman, B.; Baker, D. Computer-Based Redesignof a Protein Folding Pathway. Nat. Struct. Biol. 2001, 8,
602–605.
Janovjak, H.; Struckmeier, J.; Muller, D. J. Hydrodynamic
Eﬀects in Fast AFM Single-Molecule Force Measurements.
Eur. Biophys. J. 2005, 34, 91–96.
Liu, R. C.; Roman, M.; Yang, G. L. Correction of the Viscous
Drag Induced Errors in Macromolecular Manipulation
Experiments Using Atomic Force Microscope. Rev. Sci.
Instrum. 2010, 81, 063703.
Rico, F.; Gonzalez, L.; Casuso, I.; Puig-Vidal, M.; Scheuring, S.
High-Speed Force Spectroscopy Unfolds Titin at the Velocity of Molecular Dynamics Simulations. Science 2013, 342,
741–743.
Dufrene, Y. F.; Martinez-Martin, D.; Medalsy, I.; Alsteens, D.;
Muller, D. J. Multiparametric Imaging of Biological Systems

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