recent advances in single-molecule detection on micro- and nano-fluidic devices

Review

Recent advances in single-moleculedetection on micro- and nano-fluidic devices

Single-molecule detection (SMD) allows static and dynamic heterogeneities from

seemingly equal molecules to be revealed in the studies of molecular structures and

intra- and inter-molecular interactions. Micro- and nanometer-sized structures, including

channels, chambers, droplets, etc., in microfluidic and nanofluidic devices allow diffu-

sion-controlled reactions to be accelerated and provide high signal-to-noise ratio for

optical signals. These two active research frontiers have been combined to provide

unprecedented capabilities for chemical and biological studies. This review summarizes

the advances of SMD performed on microfluidic and nanofluidic devices published in

the past five years. The latest developments on optical SMD methods, microfluidic SMD

platforms, and on-chip SMD applications are discussed herein and future development

directions are also envisioned.

Keywords:

Lab-on-a-chip / Microfluidics / Miniaturization / Single-molecule detection /Ultra-sensitive detection DOI 10.1002/elps.201100159

1 Introduction

Since its first demonstration in 1961 [1], single-molecule

detection (SMD) has attracted significant attention in diverse

fields. Measuring signals generated by individual molecules

removes the averaging effect in classical ensemble experi-

ments. Static and dynamic heterogeneities from seemingly

equal molecules can be revealed in the studies of molecular

structures and intra- and inter-molecular interactions. Owing

to its unique advantages, SMD plays an essential role in the

investigations of chemical and biological processes.

The needs to precisely control and manipulate small

volumes of sample have led to the fast development of

microfluidics. Microfluidics has extended its widespread

usefulness into many fields and disciplines [2, 3]. Micro-

meter or nanometer-scale channels are used to efficiently

transport and analyze fluid samples. Besides the reduced

sample consumption and shortened analysis time, fast and

controllable heat supply and removal is allowed on chip,

thanks to the high surface-to-volume ratio of microchannels.

More importantly, the miniaturization of chemical reactors

(Table 1 [4]) accelerates the diffusion-controlled reactions

and dramatically increases the S/N for detection, making

microfluidics a favorable platform to provide the optimal

conditions for the analysis and manipulation of samples on

the single-molecule scale.

This review summarizes the advances in the chip-based

SMD in the past five years from the point of view of prac-

titioners of optical microscopy and spectroscopy. The read-

ers are advised to read other excellent review articles with

different focuses [4–18].

2 SMD on conventional microfluidicdevices

Laser-induced fluorescence detection has been used for

high-sensitivity analysis of minute samples in microfluidic

channels since the first demonstrations of miniaturized

devices [19, 20]. By focusing the laser beam and using a pin-

hole to block out the stray light, confocal fluorescence

microscopy provides excellent S/N for SMD in microfluidic

channels. Owing to its wide spread use and long history,

confocal fluorescence microscopy will not be discussed in

this section. Instead, we will focus on the detection schemes

that were more recently integrated with microfluidic devices.

2.1 Total internal reflection fluorescence microscopy

(TIRFM)

TIRFM has become an indispensable tool to study cellular

organization and dynamic processes that occur near the cell

Chang Liu1,2

Yueyang Qu3

Yong Luo3

Ning Fang1�

1Ames Laboratory, U.S.Department of Energy andDepartment of Chemistry, IowaState University, Ames, Iowa,USA

2Department of Chemistry,University of British Columbia,Vancouver, BC, Canada

3School of PharmaceuticalScience and Technology, DalianUniversity of Technology,Dalian, Liaoning, P. R. China

Received March 12, 2011Revised May 18, 2011Accepted June 1, 2011

Abbreviations: CICS, cylindrical illumination confocalspectroscopy; PNAs, peptide nucleic acids; SMD, singlemolecule detection; smFRET, single-molecule fluorescenceresonance energy transfer; TIRFM, total internal reflectionfluorescent microscopy

�Additional corresponding author: Ning Fang

E-mail: [email protected]

Colour Online: See the article online to view Figs. 1–3, 5 in colour.

Correspondence: Dr. Yong Luo, School of PharmaceuticalScience and Technology, Dalian University of Technology,Dalian, Liaoning, P. R. ChinaE-mail: [email protected]: 186-411-84986323

& 2011 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.electrophoresis-journal.com

Electrophoresis 2011, 32, 3308–33183308

culture and glass substrate interface [21]. It has also been

used extensively to study molecular dynamics at liquid/solid

interfaces [22–27]. All these applications rely on TIRFM’s

ability to selectively excite fluorophores very close to the

interface while minimizing background fluorescence from

out-of-focus planes. The advantages of TIRFM make it an

attractive detection scheme to be integrated with micro-

fluidic channels for studies of single molecules on or near

the channel surface.

TIRFM-based SMD experiments generally require the

use of high-magnification, high-numerical aperture (NA),

oil-immersion objective lens to collect as many photons as

possible. The working distances of these objective lenses are

typically not more than 200 mm. For objective-type TIRFM,

this is generally not a problem, because the excitation laser

beam and the emitted fluorescence photons go through the

same objective lens. However, for prism-type TIRFM, the

short working distance puts a limitation on the depth of

the microchannel and the thickness of the whole micro-

device because the excitation and emission are handled on

the opposite sides of the microdevice. Prism-type TIRFM

possesses several advantages over its objective-based coun-

terpart, such as a wider range of incident angles, higher

accuracy in incident angle determination, less scattering of

excitation light, and lower cost; therefore, much effort has

been taken to accomplish on-chip prism-type TIRFM.

The simplest solution is to create thin microfluidic

flowcells using spacers, such as double-sided tape, between

two pieces of glass coverslips [5]. Using this design, Graneli

et al. described an approach for tethering large DNA mole-

cules to the surface of a microfluidic chamber that was

rendered inert by deposition of a supported lipid bilayer, and

provided a bio-friendly environment for single-molecule

studies [28]. A nanofabricated rack pattern has also been

reported recently to align and anchor individual DNA

molecules, and the immobilized molecules could then be

visualized by TIRFM under conditions that did not require

continuous flow to stretch DNA molecules [29].

Another simple approach is to etch shallow channels

directly on the surface of glass or fused-silica coverslips. For

example, Isailovic et al. studied the affinity of DNA mole-

cules with sticky ends to the fused-silica surface of micro-

channels [30]. They showed that the affinity increased with

the number of exposed purine and pyrimidine groups,

indicating that hydrophobic forces are responsible for the

interaction of DNA molecules with the fused-silica surface.

In a more sophisticated approach, Le et al. fabricated

ultra-thin glass channels by hydrofluoric acid etching on the

coverslip while using the PDMS channels as the etching

mask. The coverslip with open microchannels was then

water bonded with another flat coverslip (Fig. 1). Single

DNA molecules (as short as 1105 base pairs) labeled with

YOYO-1 dye molecules at a ratio of 1 dye molecule per 5

base pairs were detected on the surface of this device by

prism-type TIRFM [31]. The sensitivity for imaging single

fluorescent dyes was not demonstrated in this device due to

relatively high background, which was likely caused by the

increased roughness of the etched channel surface.

The single fluorophore level detection on the prism type

TIRFM was reported on the microfluidic device recently [32].

However, the 1 mm wide straight channel in this study was

fabricated directly on the double-sided tapes, which limited

the geometry complexity and practical applications. Luo

et al. introduced a new Hi-Fiimaging microfluidic platform

and for the first time achieved the direct observation of

activation-excitation cycle in the micrometer-sized micro-

fluidic channels [33], providing the possibility of achieving

super-resolution imaging in a microfluidic device.

A useful variant of TIRFM is the so-called variable angle

epi-fluorescence or pseudo-TIRFM [34], which works at

subcritical angles that are smaller than yet still close to the

critical angle to greatly increase the illumination depth

while maintaining the low background. The Cui group used

pseudo-TRIFM to achieve background reduction for the

direct observation of axonal transport of single nerve growth

factor (NGF) in a three-compartment microfluidic chamber

[35].

2.2 Single-molecule fluorescence resonance energy

transfer (smFRET)

smFRET, a high-spatial and high-temporal resolution

method, utilizes either TIRFM or confocal fluorescence

microscopy setup for studies of structure and dynamics of

biomolecules [36]. The phenomenon of smFRET occurs over

the size range of biological macromolecules, and the

efficiency of the non-radiative energy transfer is extremely

Table 1. Scaling laws for miniaturized reaction as well as detection volume, considered for an aqueous solution of a rhodamine dye

(D 5 2.8� 10�10 m2 s�1, concentration: 1.7 nM) (from [4])

Dimension (d) 1000 mm 100 mm 10 mm 1 mm

Volume 1 mL 1 nL 1 pL 1 fL

Number of molecules 109 106 103 1

Diffusion time of a molecule over d 1.8� 106 ms 1.8� 104 ms 1.8� 102 ms 1.8 ms

Fluorescence signal per molecule 1 a.u. 1 a.u. 1 a.u. 1 a.u.

Fluorescence signal per molecule/background signal 1 a.u. 103 a.u. 106 a.u. 109 a.u.

The volume from which fluorescence is detected is assumed to be d3. Note that the fluorescent signal integrated over all molecules in

the volume per background signal remains constant, whereas the chance of detection on a single-molecule scale increases for small

detection volumes.

Electrophoresis 2011, 32, 3308–3318 Microfluidics and Miniaturization 3309


sensitive to nanoscale changes in the separation between

donor and acceptor markers. Thus, it can provide direct

information about single molecular processes and elucidate

the tertiary structure of biomolecules. The folding kinetics

of the culprit in Parkinson’s disease, protein a-synuclein,

was studied by smFRET in a rapid microfluidic mixing

platform [37]. The achieved temporal resolution was as low

as 0.2 ms. Photobleaching of fluorescent dyes is the biggest

limiting factor for acquiring high-quality smFRET signals.

To address this key limitation, Lemke et al. developed a

solution composition-independent approach, in which

deoxygenation via gas diffusion through porous PDMS

channel walls was realized to dramatically reduce oxygen-

mediated photobleaching [38]. Di Fiori and Meller published

a multimedia paper in the Journal of Visualized Experimentsto demonstrate an automated microfluidic system for long-

term smFRET measurements of surface-immobilized

biomolecules [39].

2.3 Cylindrical illumination confocal spectroscopy

(CICS)

CICS, a new variant of conventional confocal system, was

introduced for SMD on chip by Liu and Wang in 2008 [40].

In CICS, a cylindrical lens is used to shape the Gaussian

laser beam to the highly uniform 1-D sheet-like observation

volume, which results in high detection uniformity, 100%

mass detection efficiency, and high-throughput analysis

(Fig. 2). The significantly better accuracy in determining

both burst rate and burst parameters was demonstrated by

the comparison to traditional confocal measurements.

Recently, the same group achieved circulating nucleic acid

(CNA) size and quantity analysis in human serum by single-

molecule CICS on chip. The sizing analysis was performed

on serum samples from lung cancer patients at both early

and late stages without isolation or enzymatic amplification

steps, providing an alternative to the conventional PCR-

based analysis methods [41].

2.4 Other developments of detection techniques

In order to maximize photon collection efficiency in SMD,

high NA microscope objectives and precise positioning

equipments are typically necessary. An integrated optics

microfluidic device was introduced to avoid such expensive

optical parts [42]: a plano-aspheric refractive lens for

improved fluorescence excitation and a solid parabolic

reflective mirror for high-efficiency fluorescence collection.

Emory et al. presented for the first time the ability to track

single molecules simultaneously in multiple microfluidic

channels by employing a charge-coupled device (CCD)

camera operated in time-delayed integration mode as a

means of increasing throughput [43]. Chiu and co-workers

developed a method to deconvolve the number of fluor-

ophores containing fluorescence image puncta based on

single-molecule intensity distributions [44]. The methodol-

ogy was tested with the avidin-biotin-binding system for

which the predicted distribution was in good agreement

with the real experiments. Melin et al. studied key

parameters of SMD and demonstrated that the definition

of confocal volume is crucial for high-precision quantitative

measurements in microfluidic devices [45]. Zhang et al.

developed a novel approach based on single-molecule two-

color coincidence detection to evaluate the on-state quantum

dots (QDs) in a microfluidic flow [46]. Agrawal et al. used a

similar dual-color scheme to achieve real-time counting of

single native biomolecules and viruses in microfluidic

channel with the assistance of color-coded nanoparticles

[47]. Finally, silver colloid was synthesized in situ in the

microfluidic flow structure to enhance the signals from

surface enhanced Raman scattering (SERS) and surface

enhanced resonance Raman scattering (SERRS) [48].

3 SMD in integrated microfluidic devices

The integration of functional elements for sample manip-

ulation and detection provides many possibilities to achieve

high-throughput and high-sensitivity studies of complex

systems at the single-molecule level.

Figure 2. (A) Optical component diagram of CICS platform. (B)CCD image of the CICS illumination sheet focused into themicrofluidic analysis constriction. The channel boundaries aredemarcated by dashed yellow lines, while the projection of theconfocal aperture into sample space is shown in red (printedwith permission [35]).

Figure 1. Cross-sectional view of the microfluidic TIRFM-baseddevices used in [28] (printed with permission).

Electrophoresis 2011, 32, 3308–33183310 C. Liu et al.


Confinement of target molecules to small detection

volumes can effectively improve the SMD sensitivity and

increase the observation time. Kim et al. demonstrated a

microfluidic device that combined separation in microchip

capillary electrophoresis with single-molecule spectroscopy

of fluorescent biomolecules confined in sub-nanoliter

chambers formed by closing two adjacent pneumatic

microvalves [49]. In a recent study, valves and pumps were

integrated in the microfluidic mixing device, to establish the

multidimensional scanning of single molecule energy

landscapes and enzymatic activities. The miniature ring

geometry demonstrated has the volume injection errors of

0.0001% [50]. Puleo et al. used an inline micro-evaporator

cell to concentrate and transport DNA targets to the detec-

tion chamber, bridging the gap between the microliter

biological samples and the nano- to picoliter detection

volumes within the microfluidic devices [51, 52]. Single

molecules have also been shown to be confined in the

optical probe volume by hydrodynamic focusing, in which

the analyte stream was forced in the central area of the

channel, to achieve a ten-fold increase in sensitivity [53].

Besides optical trapping techniques [54, 55], the electric

or hydrodynamic field applied properly across microfluidic

channels can also trap single molecules. Cohen et al.

developed an anti-Brownian electrokinetic trap [56], in

which a time-dependent electric field was applied to induce

the electrokinetic drift to cancel the random diffusion of

single molecules in aqueous solution. Kumemura et al.

reported another trapping device using AC dielectrophoresis

[57], in which individual DNA molecules were extended and

oriented in the electric field and anchored between the

aluminum electrodes for long-term studies. The fluid field-

induced single-molecule trapping at the intersection of

microfluidic channels was reported recently [58]. A similar

study was introduced by Xu et al., and both sequence

information and restriction endonuclease cleavage kinetics

were explored on the trapped single molecule at the inter-

section [59]. Two buffer solutions flowed in the opposing

channels at the same flow rate, and single molecules were

trapped at the stagnation point at the center of the cross slot.

The kinetics of toroid formation, and the structure of DNA

inside toroids was recently investigated by utilizing both

potical and magnetic traps to manipulate single DNA

molecules in the multichannel flow cells [60].

The integration of reagent mixer with SMD has been

described in various works. Jung et al. studied single-

enzyme kinetics in a microfluidic device containing a fast

mixing element [61]. Separate substrate and enzyme

streams were rapidly mixed in the channels and encapsu-

lated in an array of individual microreactors, in which SMD

was applied to monitor the enzyme kinetics with milli-

second dead times. Ridegway et al. studied RNA–protein-

binding kinetics in an automated microfluidic reactor

capable of mixing arbitrary amounts of eight input reagents

in the reaction ring [62]. Protein folding kinetics was

investigated in a co-axial microfluidic mixer, which achieved

fast 3-D hydrodynamic focusing and rapid and efficient

diffusional mixing of the sample fluids [63]. Based on the

same principle of laminar-flow mixer, Pfeil et al. introduced

a device made of polydimethylsiloxane elastomer and fused-

silica coverglass to provide ultrasensitive detection of single

molecules under non-equilibrium conditions [64].

Multichannel microfluidic devices have been employed

to increase the throughput of on-chip analysis. Parallel

microchannels with multiple inlets and a single outlet were

shown to create a multi-laminar flow system, and each

laminar flow stream could be delivered to the desired loca-

tion in the jointing channel by varying the flow velocity

combinations for molecular patterning [65]. Okagbare et al.

demonstrated that as many as thirty parallel microchannels

can be fabricated within the large field-of-view under wide-

field illumination for the simultaneous monitoring of

single-molecule biochemical reactions [66].

While many reports were focused on integrating one

additional functional element to microdevices, multi-step

analysis of biomolecules at the single-molecule level has also

been demonstrated on micro-total analysis systems (mTAS)

by several groups. Huang et al. established an integrated

system to lyse, label, separate, and quantify the protein

contents of a single cell [67]. Nakayama et al. successfully

combined the functions of cell lysis, protein extraction,

purification, and activity assay to a single-molecule mTAS

[68]. Lam et al. developed an integrated system for the

restriction enzymatic cleavage and subsequent electro-

stretching study of freely suspended single DNA molecules

[69]. In a recent study, the extraction, manipulation and

stretching of DNA from single human chromosomes

was demonstrated on individual intergrated microfluidic

devices [70].

4 SMD in droplet-based microfluidicsystems

Droplet-based microfluidic systems have attracted much

attention in various fields [11]. The isolated compartments

within microflows have the confined volume in the pico- to

femtoliter range. Such minimized sample volume provides

a direct way of reducing fluorescent background due to

solvents and impurities [71], achieving high S/N for SMD.

The typical droplet formation process in microfluidic

channels is illustrated in Fig. 3 [11].

The encapsulation and subsequent detection of single

fluorescent molecules in microdroplets have been realized

since 1993 [72, 73], and recent developments have been

directed to improve the throughput. High-throughput

single-molecule quantification in microdroplets remained

challenging until recently. The new strategy was to employ

the CICS design by inserting a cylindrical lens into the light

path to expand the optical detection volume at the

constriction channel that forced microdroplets to squeeze

through at a slower speed [74, 75]. The entire droplet thus

could be illuminated uniformly to monitor all possible

molecular events with high temporal resolution. In another



effort, a high-throughput droplet-based single-molecule

assay for in vitro gene screening was demonstrated in an

integrated microfluidic platform [76]. In this assay, a large

number of droplets were generated and stored for hours to

allow protein expression from a single DNA template, thus

creating monoclonal droplets containing both genotype and

phenotype.

Single-molecule dynamics studies over extended time

scales require trapping droplets in the interrogation region.

Beer et al. showed a near-instantaneous droplet trapping

method using a two-position valve. The stopping time for

the droplet was as short as 38 ms [77]. Srisa-Art et al.

described that the oil flow would induce the circulation of

encapsulated molecules in the trapped droplet, enabling

multiple measurements of the same molecules over exten-

ded time periods [78].

Droplets can also be utilized indirectly for SMD. In this

approach, target molecules were not confined inside

droplets, but migrated through the submicrometer film

beneath a droplet that was closely fitted in the microchannel

[79]. The motion of single DNA molecules through the film

in the electric field was studied on this device.

5 SMD in nanofluidic devices

Besides droplets, nano-scale structure is another approach to

decrease illumination volume, and minimize the back-

ground fluorescence signals in SMD. Channels with sizes

on the order of tens to several hundreds of nanometers can

be used to constrain molecules in one or two spatial

dimensions. The polymeric properties of target molecules,

especially DNA, can be investigated within such nanos-

tructures [9, 80].

Several new approaches for fabricating nanofluidic

SMD platform have been developed in the recent years.

Liang et al. combined nanoimprint lithography with a mold

fabrication method to achieve uniform channels with

11–50 nm in width and over 1.5 cm in length [81]. The size-

controllable nanopillar arrays inside the microfluidic chan-

nels were fabricated by nanosphere lithography. The gap

distance between nanopillar could be tuned between 20 and

80 nm allowing the formation of nanofluidic system for

single DNA molecule investigation [82]. As narrow as 20 nm

channels were created by the Craighead group using the

size-reduced electrospun nanofiber method [83]. The same

group also developed the nanoglassblowing techniques for

fabricating non-planar nanofluidic platforms [84]. Recently,

Utko et al. introduced a new nanofluidic device fabrication

method with injection molding of thermoplastic polymers.

The applicability of the device was demonstrated by single

DNA molecule expriments and results were consistent with

the measurements perfomed in the costly conventional

silica nanochannels [85].

When the device scale falls below the radius of gyration

for individual molecules, the confinement will affect the

conformation of the target molecule [86]. A schematic

illustration of a nanofluidic device is shown in Fig. 4 [87].

Single polymer molecules experience a distinct force caused

by the entropic effect, which acts to move the polymer to the

area with larger configuration space [87]. When single DNA

molecules diffuse near the entrance of a nanochannel, it is

possible to force the molecules into the channels in a folded

configuration by applying an electric field. After the removal

of the electric field, the DNA molecule can extend to the

completely unfolded configuration [88]. The extension of

Figure 4. Schematic of a nanochannel device. (A) Cross-sectionof device consisting of two bonded fused-silica wafers (a, b) withthe upper one containing the structure. The microchannel wascontacted from the top of the device and fluid reservoirs (c) wereattached. Electrical connections to the channel were made byplatinum electrodes (d). (B) Close-up of the nanochannel array inthe upper wafer. DNA molecules have been drawn in the loadingzone (a), as they enter a nanochannel (b), and in an elongatedequilibrium conformation in a nanochannel (c) (printed withpermission [76]).

Figure 3. Schematic of droplet formation in a microfluidic device(printed with permission [11]).



DNA confined in nanoscale channels as a function of salt

concentration has been studied independently by two

groups [89, 90]. Different modifications to the de Gennes

theory were described to interpret the experimental data.

Nanoslits use one nanoscale dimension (depth) to

restrict the movement of DNA molecules to a 2-D plane. The

effects of confinement on electrophoretic stretching of

single DNA molecules were studied by Balducci et al. [91].

Reisner et al. constructed embedded nanopit arrays and

showed that the position and conformation of individual

DNA molecules could be well controlled in the nanoslits by

adjusting the spacing, organization, and placement of

nanopits [92].

Besides the manipulation of single DNA molecules,

SMD in nanochannels were reported in other fields. High-

throughput single-molecule epigenetic analysis was estab-

lished for the first time by Cipriany et al. in nanofluidic

channels [93]. Time-coincident fluorescent signatures of

both DNAs and histone proteins were analyzed to identify

the methylation on individual molecules. Fluorescence

signals from single partially denatured DNA molecules were

reported as a sequence-depended ‘‘barcode’’ with high

sensitivity to the sequence variation, suitable for investigat-

ing the long-range structure of entire genomes [94]. Yama-

moto et al. combined SMD with an identification element

and a low path switch to build a nanofluidic single-molecule

sorter [95].

In-channel nanopores are another important platform

for investigating dynamics and thermodynamics of confined

single molecules. Shiu et al. measured the lengths of stret-

ched DNA molecules in the ordered in-channel nanopores

[96]. The channel current through membrane-bound ion

channels on reconstituted lipid bilayer was recorded by

Suzuki et al. [97, 98] and Hromada et al. [99]. This technique

can be used for studying electrophysiology at the single-

molecule level. The simultaneous monitoring of multiple

ionic currents through self-assembled bilayer membrane

was reported as well in a microarray system [100].

6 Applications of micro- and nano-fluidicsfeaturing SMD

A tremendous amount of effort has been put into

biophysical and bioanalytical studies of single macromole-

cules in micro- or nano-fluidic devices. Many applications

have been discussed together with the corresponding

microfluidic devices in the previous sections. In this section,

we will summarize other interesting and relevant applica-

tions.

6.1 DNA manipulation

To investigate DNA–protein interactions at the single-

molecule level, individual DNA molecules are often

immobilized on the surface to suppress Brownian motion,

so that the shape and position of the DNA, as well as protein

motion along the DNA contour, can be followed precisely

[101]. At slightly acidic conditions (pH 5.5–6.6), the

molecular combing technique can readily immobilize

DNA molecules onto a hydrophobic microchannel surface

[102]. In order to work at the physiological pH 7.4 and to

avoid the overstretching of DNA molecules, Dukkipati et al.

introduced a protein-assisted DNA immobilization and

stretching approach, in which the proteins on the

DNA–protein complex adsorbed on the channel surface to

stretch the DNA under a hydrodynamic flow [103]. In

another study, the surface of microchannel was rendered

passive to adsorption by functionalizing with a block

copolymer, poly(L-lysine-graft-polyethylene glycol), thus

minimizing the fluorescence background from the non-

targeted adsorbed molecules, and more importantly, the

channel surface properties were stable under either wet or

dry conditions [104].

Manipulation of DNA molecules has also been

demonstrated in free solution without immobilization. The

elongation and migration of single DNA molecules in

oscillatory shear flows were characterized systematically as a

function of molecular size, shear rate, and oscillatory

frequency [105]. The pressure-driven oscillatory shear flows

stretched DNA molecules and forced them to migrate away

from the walls and focus in the channel centerline [105].

Free DNA molecules can also be stretched in sudden mixed

shear and elongational microflows formed at different

funnel-shaped microfluidic devices [106]. In a more recent

study, DNA stretching was used to analysis viruses and

toxins in an integrated microfluidic system [107]. Fast

transition between the stretching and compression of the

DNA was observed in an electric field gradient generated in

tapered contraction–expansion microchannels [108]. Precise

control over the lateral position and motion of DNA mole-

cules in sub-micrometer channels was demonstrated by

adjusting the amplitude or frequency of the AC potential

applied to the electrode pair on both sides of the channels

[109].

Manipulation of DNA molecules in polymer solutions

and gels has been reported, too. Polymer solutions have long

been studied for their interesting nonlinear viscoelastic

response properties and played an important role in the field

of electrophoretic DNA separations. Electrophoretic DNA

migration modalities above and below entanglement were

both demonstrated [110]. A cross-linked gel patterned inside

in a hyperbolic contraction microchannel was shown to

stretch DNA molecules while they reptated through the gel

under the influence of electric field gradients [111].

6.2 DNA sequencing

Single-molecule sequencing provides several advantages

over the conventional sequencing methods. It eliminates

the need for cloning to result in increased throughput and

reduced costs. In addition, it promises the reading length of



tens of kilobases. Pacific Biosciences reported a real-time

single-molecule DNA sequencing system, in which a single

DNA polymerase molecule is confined together with the

DNA in a zero-mode waveguide. The temporal sequence of

distinguishable labeled dNTPs adding onto the polymeriz-

ing chain was recorded to achieve DNA sequencing [112].

Bashford et al. introduced an automated single-molecule

DNA sequencing system, in which DNA molecules were

immobilized on pressure-trapped beads and then physically

extended into a downstream channel under electric field

[113]. In another effort by Dylla-Spears et al. [58], DNA

molecules were trapped at a stagnation point at the center of

a planar extensional flow and elongated along the outflow

axis in the hydrodynamic fluid field, and both DNA

backbone and the markers bound along the stretched

DNA could be observed directly using fluorescence micro-

scopy.

Peptide nucleic acids (PNAs) are synthetic molecules

composed of a peptide backbone and nucleic acid bases, and

they have several advantages for single-molecule level DNA

studies. PNAs are compatible with cofactors and non-

destructive to double-strand DNA and do not require liga-

tion and bind with high specificity and yield [114]. Deter-

mining target DNA sequences with labeled PNAs was

shown to be comparable to other single-molecule sequen-

cing methods [115, 116].

6.3 Conformation probing

High-spatial resolution of SMD, especially in smFRET,

makes it a powerful tool for studies of single-molecule

conformation changes. The individual RNA molecules were

immobilized on the surface of microfluidic flow chamber

and probed by smFRET to reveal the RNA conformation

change accompanying Mg21 concentration jump [117]. The

smFRET study on protein folding in chaperonin cavities

showed that the confinement in the chaperonin decelerated

the folding of C-terminal domain of the protein, while the

folding rate of the N-terminal domain was unaffected [118].

The same group used the combination of smFRET and

kinetic synchrotron radiation circular dichroism experi-

ments to probe the conformational ensemble of the

collapsed unfolded state of a protein under near native

conditions in microfluidic devices, and successfully differ-

entiated the folded and unfolded subpopulations at equili-

brium and provided information on long-range

intramolecular distance distributions [119].

6.4 Protein interactions

Conventional techniques for studying protein interactions,

e.g. 2-D electrophoresis-mass spectrometry and immuno-

precipitation/Western blot, usually require larger sample

size and longer processing time than the SMD-based

techniques. Single-molecule studies of protein-protein

interactions provide more insightful information on

dynamics. Chou et al. developed a digital protein analysis

platform to measure the interaction ratio between oncogenic

signaling protein complexes before and after ligand

stimulation [120]. The event of RAD51 proteins disassem-

bling from nucleoprotein filaments under external tension

was studied in a multichannel microfluidic flow cell [121].

Arakawa et al. studied single protein-binding and dissocia-

tion processes using a microfluidic TIRFM imaging system

integrated with a rapid switching microvalve system for

multi-reagent exchange [122].

Protein–protein interactions were also used to detect

individual molecules in microfluidic channels. Staphylo-

coccal Enterotoxin B (SEB) was detected at the single-

molecule level using a mobile sandwich immunoassay on

gliding microtubules that were modified postpolymerization

with anti-SEB [123]. Nalefski et al. developed a microfluidic

system enabling single-molecule protein detection and

quantification in heterogeneous immunoassays [124].

6.5 On-chip SMD in cells

SMD in living cells is still challenging due to high cellular

autofluorescence background and photobleaching at high

intracellular oxygen concentrations. Cai et al. successfully

demonstrated the real-time observation of stochastic expres-

sion of a-galactosidase at the single-molecule level in living

Escherichia coli cells and studied the key parameters for

protein expression, including burst size and frequency [125].

Huang et al. introduced a microfluidic device for counting

multiple low-copy number proteins in a single living cell

that cannot be distinguished by fluorescence properties

(Fig. 5) [126]. This system was utilized to analyze

phycobiliprotein contents in individual cyanobacterial cells

and observed marked differences in the levels of specific

complexes in the cells grown under different nitrogen levels.

6.6 Other applications

Single-molecule imaging provides a practical approach for

generating molecular pictures of flow profiles in real time.

Fluorescence correlation spectroscopy (FCS) provides the

ability to probe flow profiles and mass transport in

microfluidic channels without mechanically accessing the

channels. Single-molecule FCS was used to map the vortex-

like hydrodynamic flow at a T-shaped junction of micro-

channels [127], and to acquire direct kinetics measurements

in a rapid continuous flow mixer using zero-mode

waveguides [128]. A modified surface forces apparatus for

single-molecule tracking and a number of data analysis

methods were introduced for mapping flow profiles in shear

flows of ultrathin liquid films [129–131].

Temperature measurement in fluids relies traditionally

on the intensity variation determination for probe molecules

with temperature-dependent quantum yields [132].



However, local dye concentration and background fluores-

cence can also affect signal intensity, leading to less quan-

titative information on the absolute temperature. Based on

SMD in channels, a novel molecular thermometer was

developed by utilizing enhanced green fluorescent protein

(EGFP) as the probe molecule, because the relaxation time

of EGFP is highly temperature sensitive [133].

7 Concluding remarks and outlook

In this review, we survey the advances in the micro-and-

nano-fluidics-based SMD. DNA has been the most intensely

studied target in microfluidic devices at the single-molecule

level, which is not by accident. Not only is the under-

standing of biochemistry and biophysics of DNA essential to

the understanding of living organisms and the search for

cures for disease, but also are there fewer technical obstacles

to overcome for visualizing single DNA molecules. The size

of DNA and its structure of being a long, linear polymer

make it convenient to label hundreds of fluorescent dyes on

a single DNA molecule. The strong fluorescence signal is

much needed in dealing with the higher background for

SMD in microchannels. As researchers work more

frequently with smaller and dimmer single biomolecules,

optically perfect microchannels become more and more

important. All surfaces of optically transparent building

blocks in the optical path should be flat and parallel to each

other to reduce background noise and minimize inter-

ference fringes. The scattering effect at channel walls and

other boundaries may be reduced or even eliminated by

matching the refractive indices of channel materials to the

fluid inside the microchannels.

Recent breakthroughs in lens-based optical microscopy

have overcome the diffraction barrier imposed by the wave

nature of light [134]. For example, stochastic optical recon-

struction microscopy (STORM) [135–137], as well as photo-

activated localization microscopy (PALM) [138, 139], realizes

the high resolution by super-localizing randomly activated

photo-switchable fluorophores. To perform STORM or

PALM, the imaging platform should be able to detect single

fluorophores and the resolution depends on the photon

collection efficiency. We expect these novel imaging tools to

be incorporated into the designs of microfluidic devices to

provide unprecedented capabilities of super-resolution

imaging under external perturbation on biological samples.

Another area expecting great efforts is to fully integrate

SMD techniques into microdevices to decrease reliance on

external equipment and achieve higher degree of portability.

Historically, the SMD experiments were limited to a small

number of laboratories equipped with lasers, sensitive

detectors, and other expensive electronic devices. The costs

Figure 5. Single-cell analysischip. (A) Layout of the single-cell chip, showing the cell-manipulation section on theleft and the molecule-count-ing section on the right. (B)Analysis procedure for amammalian or insect cell. (C)Schematic illustration of theexcitation laser focused by themicroscope objective and thedimensions of the molecule-counting channel. (D) Oneframe from the CCD imagesof fluorescent molecules flow-ing across the molecule-counting section (upperpanel) and the identificationresults (lower panel) (printedwith permission [115]).



of running SMD experiments have come down dramatically

in the past decade as all of these essential components

became affordable to most laboratories. Further integration

of SMD and micro- and nano-fluidics may result in

commercial products for ultrahigh-sensitivity clinical and

pharmaceutical applications.

This work was supported by the ‘‘Fundamental ResearchFunds for the Central Universities, China’’ (DUT10RC(3)92and DUT11SM11)and U.S. Department of Energy, Office ofBasic Energy Sciences, Division of Chemical Sciences, Geos-ciences, and Biosciences through the Ames Laboratory. The AmesLaboratory is operated for the U.S. Department of Energy byIowa State University under contract no. DE-AC02-07CH11358. The authors specially thank Prof. David Chen ofUniversity of British Columbia for his financial support forChang Liu’s visit to Ames Lab.

The authors have declared no conflict of interest.

8 References

[1] Rotman, B., Proc. Natl. Acad. Sci. USA 1961, 47,1981–1991.

[2] Arora, A., Simone, G., Salieb-Beugelaar, G. B., Kim, J.T., Manz, A., Anal. Chem. 2010, 82, 4830–4847.

[3] Salieb-Beugelaar, G. B., Simone, G., Arora, A., Philippi,A., Manz, A., Anal. Chem. 2010, 82, 4848–4864.

[4] Dittrich, P. S., Manz, A., Anal. Bioanal. Chem. 2005,382, 1771–1782.

[5] Brewer, L. R., Bianco, P. R., Nat. Methods 2008, 5,517–525.

[6] Chiu, D. T., Anal. Bioanal. Chem. 2007, 387, 17–20.

[7] Craighead, H., Nature 2006, 442, 387–393.

[8] Leuba, S. H., Wheeler, T. B., Cheng, C.-M., Le Duc, P. R.,Fernandez-Sierra, M., Quinones, E., Methods 2009, 47,214–222.

[9] Levy, S. L., Craighead, H. G., Chem. Soc. Rev. 2010, 39,1133–1152.

[10] Spetzler, D., York, J., Dobbin, C., Martin, J., Ishmu-khametov, R., Day, L., Yu, J., Kang, H., Porter, K.,Hornung, T., Frasch, W. D., Lab Chip 2007, 7,1633–1643.

[11] Theberge, A. B., Courtois, F., Schaerli, Y., Fischlechner,M., Abell, C., Hollfelder, F., Huck, W. T. S., Angew.Chem. – Int. Edit. 2010, 49, 5846–5868.

[12] Varghese, S. S., Zhu, Y. G., Davis, T. J., Trowell, S. C.,Lab Chip 2009, 10, 1355–1364.

[13] Wang, T.-H., Puleo, C. M., Yeh, H.-C., Integr. BiochipsDNA Anal. 2007, 139–150.

[14] Witek, M. A., Hupert, M. L., Soper, S. A., Bio-MEMS2007, 391–441.

[15] Zhang, C., Xing, D., Chem. Rev. 2010, 110, 4910–4947.

[16] Erickson, D., Serey, X., Chen, Y. F., Mandal, S., LabChip 2011, 11, 995–1009.

[17] Yu, T. A., Martinez, M. M., Pappas, D., Appl. Spec.

2011, 65, 115A–124A.

[18] Yin, Y., Qiu, T., Zhang, W. J., Chu, P. K., J. Mater. Res.

2011, 26, 170–185.

[19] Harrison, D. J., Fluri, K., Seiler, K., Fan, Z. H.,

Effenhauser, C. S., Manz, A., Science 1993, 261,895–897.

[20] Harrison, D. J., Manz, A., Fan, Z. H., Ludi, H., Widmer,

H. M., Anal. Chem. 1992, 64, 1926–1932.

[21] Axelrod, D., in: Correia, J. J., Detrich, H. W. (Eds.),

Biophysical Tools for Biologists, Volume Two: In VivoTechniques, Academic Press, Boston, MA 2008, pp.169–221.

[22] Xu, X. H., Yeung, E. S., Science 1997, 275, 1106–1109.

[23] Xu, X. H. N., Yeung, E. S., Science 1998, 281,

1650–1653.

[24] Kang, S. H., Shortreed, M. R., Yeung, E. S., Anal. Chem.

2001, 73, 1091–1099.

[25] Kang, S. H., Yeung, E. S., Anal. Chem. 2002, 74,

6334–6339.

[26] He, Y., Li, H. W., Yeung, E. S., J. Phys. Chem. B 2005,

109, 8820–8832.

[27] Fang, N., Zhang, H., Li, J. W., Li, H. W., Yeung, E. S.,

Anal. Chem. 2007, 79, 6047–6054.

[28] Graneli, A., Yeykal, C. C., Prasad, T. K., Greene, E. C.,

Langmuir 2006, 22, 292–299.

[29] Gorman, J., Fazio, T., Wang, F., Wind, S., Greene, E. C.,

Langmuir 2010, 26, 1372–1379.

[30] Isailovic, S., Li, H. W., Yeung, E. S., J. Chromatogr. A

2007, 1150, 259–266.

[31] Le, N. C. H., Yokokawa, R., Dao, D. V., Nguyen, T. D.,

Wells, J. C., Sugiyama, S., Lab Chip 2009, 9, 244–250.

[32] Wang, B., Ho, J., Fei, J. Y., Gonzalez, R. L., Lin, Q. A.,

Lab Chip 2011, 11, 274–281.

[33] Luo, Y., Sun, W., Liu, C., Wang, G. F., Fang, N., Anal.

Chem. 2011, 83, 5073–5077.

[34] Konopka, C. A., Bednarek, S. Y., Plant J. 2008, 53,

186–196.

[35] Zhang, K., Osakada, Y., Vrljic, M., Chen, L., Mudrakola,

H. V., Cui, B., Lab Chip 2010, 10, 2566–2573.

[36] Varghese, S. S., Zhu, Y. G., Davis, T. J., Trowell, S. C.,

Lab Chip 2009, 10, 1355–1364.

[37] Gambin, Y., VanDelinder, V., Ferreon, A. C. M., Lemke,

E. A., Groisman, A., Deniz, A. A., Nat. Methods 2011, 8,239–U277.

[38] Lemke, E. A., Gambin, Y., Vandelinder, V., Brustad, E.

M., Liu, H.-W., Schultz, P. G., Groisman, A., Deniz, A.A., J. Am. Chem. Soc. 2009, 131, 13610–13612.

[39] Di Fiori, N., Meller, A., J. Vis. Exp. 2009, e1542.

[40] Liu, K. J., Wang, T.-H., Biophys. J. 2008, 95,

2964–2975.

[41] Liu, K. J., Brock, M. V., Shih, I.-M., Wang, T.-H., J. Am.

Chem. Soc. 2010, 132, 5793–5798.

[42] Krogmeier, J. R., Schaefer, I., Seward, G., Yantz, G. R.,

Larson, J. W., Lab Chip 2007, 7, 1767–1774.

[43] Emory, J. M., Soper, S. A., Anal. Chem. 2008, 80,

3897–3903.



[44] Mutch, S. A., Fujimoto, B. S., Kuyper, C. L., Kuo, J. S.,Bajjalieh, S. M., Chiu, D. T., Biophys. J. 2007, 92,2926–2943.

[45] Melin, J., Jarvius, J., Goeransson, J., Nilsson, M., Anal.Biochem. 2007, 368, 230–238.

[46] Zhang, C.-Y., Yang, K., Anal. Bioanal. Chem. 2010, 397,703–708.

[47] Agrawal, A., Zhang, C., Byassee, T., Tripp, R. A., Nie,S., Anal. Chem. 2006, 78, 1061–1070.

[48] Wilson, R., Bowden, S. A., Parnell, J., Cooper, J. M.,Anal. Chem. 2010, 82, 2119–2123.

[49] Kim, S., Huang, B., Zare, R. N., Lab Chip 2007, 7,1663–1665.

[50] Kim, S., Streets, A. M., Lin, R. R., Quake, S. R., Weiss,S., Majumdar, D. S., Nat. Methods 2011, 8,242–283.

[51] Puleo, C. M., Wang, T.-H., Lab Chip 2009, 9, 1065–1072.

[52] Puleo, C. M., Yeh, H. C., Liu, K. J., Rane, T., Wang, T. H.,21st IEEE International Conference on Micro ElectroMechanical Systems, Technical Digest, 2008, 1,200–203.

[53] de Mello, A. J., Edel, J. B., J. Appl. Phys. 2007, 101,084903.

[54] Schmidt, B. S., Yang, A. H. J., Erickson, D., Lipson, M.,Opt. Express 2007, 15, 14322–14334.

[55] Yang, A. H. J., Moore, S. D., Schmidt, B. S., Klug, M.,Lipson, M., Erickson, D., Nature 2009, 457, 71–75.

[56] Cohen, A. E., Moerner, W. E., Opt. Express 2008, 16,6941–6956.

[57] Kumemura, M., Collard, D., Yamahata, C., Sakaki, N.,Hashiguchi, G., Fujita, H., ChemPhysChem 2007, 8,1875–1880.

[58] Dylla-Spears, R., Townsend, J. E., Jen-Jacobson, L.,Sohn, L. L., Muller, S. J., Lab Chip 2010, 10, 1543–1549.

[59] Xu, W. L., Muller, S. J., Lab Chip 2011, 11, 435–442.

[60] Brewer, L. R., Integr. Biol. 2011, 3, 540–547.

[61] Jung, S.-Y., Liu, Y., Collier, C. P., Langmuir 2008, 24,4439–4442.

[62] Ridgeway, W. K., Seitaridou, E., Phillips, R., William-son, J. R., Nucleic Acids Res. 2009, 37, e142.

[63] Hamadani, K. M., Weiss, S., Proceedings of SPIE, 2008,6862, 68620A.

[64] Pfeil, S. H., Wickersham, C. E., Hoffmann, A., Lipman,E. A., Rev. Sci. Instrum. 2009, 80, 055105.

[65] Lee, S. W., Yamamoto, T., Noji, H., Fujii, T., EnzymeMicrob. Technol. 2006, 39, 519–525.

[66] Okagbare, P. I., Soper, S. A., Analyst 2009, 134, 97–106.

[67] Huang, B., Wu, H. K., Bhaya, D., Grossman, A., Granier,S., Kobilka, B. K., Zare, R. N., Science 2007, 315, 81–84.

[68] Nakayama, T., Namura, M., Tabata, K. V., Noji, H.,Yokokawa, R., Lab Chip 2009, 9, 3567–3573.

[69] Lam, L., Sakakihara, S., Ishizuka, K., Takeuchi, S., Noji,H., Lab Chip 2007, 7, 1738–1745.

[70] Rasmussen, K. H., Marie, R., Lange, J. M., Svendsen,W. E., Kristensen, A., Mir, K. U., Lab Chip 2011, 11,1431–1433.

[71] de Mello, A. J., Lab Chip 2003, 3, 29N–34N.

[72] Barnes, M. D., Lermer, N., Kung, C. K., Whitten, W. B.,Ramsey, J. M., Hill, S. C., Opt. Lett. 1997, 22,1265–1267.

[73] Barnes, M. D., Ng, K. C., Whitten, W. B., Ramsey, J. M.,Anal. Chem. 1993, 65, 2360–2365.

[74] Srisa-Art, M., de Mello, A. J., Edel, J. B., Chem.Commun. 2009, 6548–6550.

[75] Rane, T. D., Puleo, C. M., Liu, K. J., Zhang, Y., Lee, A. P.,Wang, T. H., Lab Chip 2010, 10, 161–164.

[76] Courtois, F., Olguin, L. F., Whyte, G., Bratton, D., Huck,W. T. S., Abell, C., Hollfelder, F., ChemBioChem 2008,9, 439–446.

[77] Beer, N. R., Rose, K. A., Kennedy, I. M., Lab Chip 2009,9, 841–844.

[78] Srisa-Art, M., de Mello, A. J., Edel, J. B., J. Phys. Chem.B 2010, 114, 15766–15772.

[79] Hsieh, S.-F., Wei, H.-H., Phys. Rev. E 2009, 79, 021901.

[80] Mannion, J. T., Craighead, H. G., Biopolymers 2007, 85,131–143.

[81] Liang, X. G., Morton, K. J., Austin, R. H., Chou, S. Y.,Nano Lett. 2007, 7, 3774–3780.

[82] Kuo, C.-W., Wei, K. H., Lin, C.-H., Shiu, J.-Y., Chen, P.,Electrophoresis 2008, 29, 2931–2938.

[83] Park, S.-m., Huh, Y. S., Szeto, K., Joe, D. J., Kameoka,J., Coates, G. W., Edel, J. B., Erickson, D., Craighead, H.G., Small 2010, 6, 2420–2426.

[84] Strychalski, E. A., Stavis, S. M., Craighead, H. G.,Nanotechnology 2008, 19, 315301.

[85] Utko, P., Persson, F., Kristensen, A., Larsen, N. B., LabChip 2011, 11, 303–308.

[86] Tegenfeldt, J. O., Prinz, C., Cao, H., Chou, S., Reisner,W. W., Riehn, R., Wang, Y. M., Cox, E. C., Sturm, J. C.,Silberzan, P., Austin, R. H., Proc. Natl. Acad. Sci. USA2004, 101, 10979–10983.

[87] Mannion, J. T., Reccius, C. H., Cross, J. D., Craighead,H. G., Biophys. J. 2006, 90, 4538–4545.

[88] Levy, S. L., Mannion, J. T., Cheng, J., Reccius, C. H.,Craighead, H. G., Nano Lett. 2008, 8, 3839–3844.

[89] Reisner, W., Beech, J. P., Larsen, N. B., Flyvbjerg, H.,Kristensen, A., Tegenfeldt, J. O., Phys. Rev. Lett. 2007,99, 058302.

[90] Zhang, C., Zhang, F., van Kan, J. A., van der Maarel, J.R. C., J. Chem. Phys. 2008, 128, 225109.

[91] Balducci, A. G., Tang, J., Doyle, P. S., Macromolecules2008, 41, 9914–9918.

[92] Reisner, W., Larsen, N. B., Flyvbjerg, H., Tegenfeldt, J.O., Kristensen, A., Proc. Natl. Acad. Sci. USA 2009, 106,79–84.

[93] Cipriany, B. R., Zhao, R., Murphy, P. J., Levy, S. L., Tan,C. P., Craighead, H. G., Soloway, P. D., Anal. Chem.2010, 82, 2480–2487.

[94] Reisner, W., Larsen, N. B., Silahtaroglu, A., Kristensen,A., Tommerup, N., Tegenfeldt, J. O., Flyvbjerg, H.,Proc. Natl. Acad. Sci. USA 2010, 107, 13294–13299.

[95] Yamamoto, T., Fujii, T., Nanotechnology 2010, 21,395502.

[96] Shiu, J.-Y., Whang, W.-T., Chen, P., J. Chromatogr. A2008, 1206, 72–76.



[97] Suzuki, H., Tabata, K. V., Noji, H., Takeuchi, S., Lang-muir 2006, 22, 1937-1942.

[98] Suzuki, H., Tabata, K. V., Noji, H., Takeuchi, S.,Biosensors Bioelectronics 2007, 22, 1111–1115.

[99] Hromada, L. P., Nablo, B. J., Kasianowicz, J. J., Gaitan,M. A., DeVoe, D. L., Lab Chip 2008, 8, 602–608.

[100] Osaki, T., Suzuki, H., Le Pioufle, B., Takeuchi, S., Anal.Chem. 2009, 81, 9866–9870.

[101] van Mameren, J., Peterman, E. J. G., Wuite, G. J. L.,Nucleic Acids Res. 2008, 36, 4381–4389.

[102] Petit, C. A. P., Carbeck, J. D., Nano Lett. 2003, 3,1141–1146.

[103] Dukkipati, V. R., Kim, J. H., Pang, S. W., Larson, R. G.,Nano Lett. 2006, 6, 2499–2504.

[104] Vasdekis, A. E., O’Neil, C. P., Hubbell, J. A., Psaltis, D.,Biomacromolecules 2010, 11, 827–831.

[105] Jo, K., Chen, Y.-L., de Pablo, J. J., Schwartz, D. C., LabChip 2009, 9, 2348–2355.

[106] Larson, J. W., Yantz, G. R., Zhong, Q., Charnas, R.,D’Antoni, C. M., Gallo, M. V., Gillis, K. A., Neely, L. A.,Phillips, K. M., Wong, G. G., Gullans, S. R., Gilmanshin,R., Lab Chip 2006, 6, 1187–1199.

[107] Meltzer, R. H., Krogmeier, J. R., Kwok, L. W., Allen, R.,Crane, B., Griffis, J. W., Knaian, L., Kojanian, N.,Malkin, G., Nahas, M. K., Papkov, V., Shaikh, S.,Vyavahare, K., Zhong, Q., Zhou, Y., Larson, J. W.,Gilmanshin, R., Lab Chip 2011, 11, 863–873.

[108] Hu, X., Wang, S., Lee, L. J., Phys. Rev. E 2009, 79, 041911.

[109] Cardozo, B. L., Pang, S. W., J. Vacuum Sci. Technol. B:Microelectronics Nanometer Struc. – Process.,Measur. Phenomena 2008, 26, 2578–2582.

[110] Chiesl, T. N., Forster, R. E., Root, B. E., Larkin, M.,Barron, A. E., Anal. Chem. 2007, 79, 7740–7747.

[111] Randall, G. C., Schultz, K. M., Doyle, P. S., Lab Chip2006, 6, 516–525.

[112] Eid, J., Fehr, A., Gray, J., Luong, K., Lyle, J., Otto, G.,Peluso, P., Rank, D., Baybayan, P., Bettman, B., Bibillo,A., Bjornson, K., Chaudhuri, B., Christians, F., Cicero,R., Clark, S., Dalal, R., Dewinter, A., Dixon, J., Foquet,M., Gaertner, A., Hardenbol, P., Heiner, C., Hester, K.,Holden, D., Kearns, G., Kong, X. X., Kuse, R., Lacroix,Y., Lin, S., Lundquist, P., Ma, C. C., Marks, P., Maxham,M., Murphy, D., Park, I., Pham, T., Phillips, M., Roy, J.,Sebra, R., Shen, G., Sorenson, J., Tomaney, A.,Travers, K., Trulson, M., Vieceli, J., Wegener, J., Wu,D., Yang, A., Zaccarin, D., Zhao, P., Zhong, F., Korlach,J., Turner, S., Science 2009, 323, 133–138.

[113] Bashford, G., Lamb, D., Grone, D., Eckles, B., Kornel-sen, K., Middendorf, L., Williams, J., Opt. Express 2008,16, 3445–3455.

[114] Chan, E. Y., Goncalves, N. M., Haeusler, R. A., Hatch, A.J., Larson, J. W., Maletta, A. M., Yantz, G. R., Carstea,E. D., Fuchs, M., Wong, G. G., Gullans, S. R., Gilman-shin, R., Genome Res. 2004, 14, 1137–1146.

[115] Zohar, H., Hetherington, C. L., Bustamante, C., Muller,S. J., Nano Lett. 2010, 10, 4697–4701.

[116] Protozanova, E., Zhang, M., White, E. J., Mollova, E. T.,Ten Broeck, D., Fridrikh, S. V., Cameron, D. B.,Gilmanshin, R., Anal. Biochem. 2010, 402, 83–90.

[117] Qu, X. H., Smith, G. J., Lee, K. T., Sosnick, T. R., Pan, T.,Scherer, N. F., Proc. Natl. Acad. Sci. USA 2008, 105,6602–6607.

[118] Hofmann, H., Hillger, F., Pfeil, S. H., Hoffmann, A.,Streich, D., Haenni, D., Nettels, D., Lipman, E. A.,Schuler, B., Proc. Natl. Acad. Sci. USA 2010, 107,11793–11798.

[119] Hoffmann, A., Kane, A., Nettls, D., Hertzog, D. E.,Baumgartel, P., Lengefeld, J., Reichardt, G., Horsley, D.A., Seckler, R., Bakajin, O., Schuler, B., Proc. Natl.Acad. Sci. USA 2007, 104, 105–110.

[120] Chou, C.-K., Jing, N., Yamaguchi, H., Tsou, P.-H., Lee,H.-H., Chen, C.-T., Wang, Y.-N., Hong, S., Su, C.,Kameoka, J., Hung, M.-C., Lab Chip 2010, 10,1793–1798.

[121] van Mameren, J., Modesti, M., Kanaar, R., Wyman, C.,Peterman, E. J. G., Wuite, G. J. L., Nature 2009, 457,745–748.

[122] Arakawa, T., Sameshima, T., Sato, Y., Ueno, T., Shir-asaki, Y., Funatsu, T., Shoji, S., Sensors Actuators, B:Chem. 2007, B128, 218–225.

[123] Soto, C. M., Martin, B. D., Sapsford, K. E., Blum, A. S.,Ratna, B. R., Anal. Chem. 2008, 80, 5433–5440.

[124] Nalefski, E. A., D’Antoni, C. M., Ferrell, E. P., Lloyd, J.A., Qiu, H., Harris, J. L., Whitney, D. H., Clin. Chem.2006, 52, 2172–2175.

[125] Cai, L., Friedman, N., Xie, X. S., Nature 2006, 440,358–362.

[126] Huang, B., Wu, H., Bhaya, D., Grossman, A., Granier,S., Kobilka, B. K., Zare, R. N., Science 2007, 315, 81–84.

[127] Liu, K., Tian, Y., Burrows, S. M., Reif, R. D., Pappas, D.,Anal. Chim. Acta 2009, 651, 85–90.

[128] Liao, D., Galajda, P., Riehn, R., Ilic, R., Puchalla, J. L.,Yu, H. G., Craighead, H. G., Austin, R. H., Opt. Express2008, 16, 10077–10090.

[129] Schob, A., Pumpa, M., Selmke, M., Cichos, F., J. Phys.Chem. C 2010, 114, 4479–4485.

[130] Schob, A., Cichos, F., J. Phys. Chem. B 2006, 110,4354–4358.

[131] Schob, A., Cichos, F., Chem. Phys. Lett. 2010, 484,192–196.

[132] Karstens, T., Kobs, K., J. Phys. Chem. 1980, 84,1871–1872.

[133] Wong, F. H. C., Banks, D. S., Abu-Arish, A., Fradin, C.,J. Am. Chem. Soc. 2007, 129, 10302–10303.

[134] Hell, S. W., Nat. Methods 2009, 6, 24–32.

[135] Huang, B., Wang, W. Q., Bates, M., Zhuang, X. W.,Science 2008, 319, 810–813.

[136] Bates, M., Huang, B., Dempsey, G. T., Zhuang, X. W.,Science 2007, 317, 1749–1753.

[137] Rust, M. J., Bates, M., Zhuang, X. W., Nat. Methods2006, 3, 793–795.

[138] Betzig, E., Patterson, G. H., Sougrat, R., Lindwasser, O.W., Olenych, S., Bonifacino, J. S., Davidson, M. W.,Lippincott-Schwartz, J., Hess, H. F., Science 2006, 313,1642–1645.

[139] Hess, S. T., Girirajan, T. P. K., Mason, M. D., Biophys. J.2006, 91, 4258–4272.



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