2267
Electrophoresis 2014, 35, 2267–2278
Chaoqing Dong Jicun Ren State Key Laboratory of Metal Matrix Composites, School of Chemistry & Chemical Engineering, Shanghai Jiao Tong University, Shanghai, P. R. China
Received December 24, 2013 Revised March 10, 2014 Accepted March 21, 2014
Review
Coupling of fluorescence correlation spectroscopy with capillary and microchannel analytical systems and its applications In the past decade, fluorescence correlation spectroscopy (FCS) has become an ultrasensitive and noninvasive single-molecule detection technique, which is widely applied in the physical, chemical, and life-science research. The coupling of FCS with narrow channel flow systems including the ones based on capillary provide the important, convenient, and sensitive assay platforms for probing and understanding the behavior of single molecules or nanoparticles with improved temporal and spatial resolution and need for less sample volume among other advantages. This review focus on different approaches for FCS with capillary and microchannel analytical systems and its applications in confined diffusion study, flow profiles, and imaging of narrow channel, multicomponent analyses such as protein, DNA analysis, and characterization on nanoparticles. Keywords: Capillary electrophoresis / Coupling / Fluorescence correlation spectroscopy / TAS DOI 10.1002/elps.201300648
1 Introduction Fluorescence correlation spectroscopy (FCS) is an ultrasensitive and noninvasive technique for single-molecule detection that uses statistical analysis of the fluorescence fluctuation emitted from a small and optically well-defined open volume element [1–3]. In contrast to other fluorescence techniques, FCS can provide not only the emission intensity of fluorophors in the volume, but also other important parameters such as the diffusion or mobility coefficients of fluorophors, etc. because of the minor intensity fluctuations caused by the minute deviations of molecules from thermal equilibrium. Thus, any factors that lead to the deviations from equilibrium can be investigated by FCS including diffusion, physical or chemical reactions, aggregation, etc. Using FCS, local concentration, diffusion coefficients, chemical reaction rate as well as photophysical properties of fluorophors can be determined.
Correspondence: Professor Jicun Ren, State Key Laboratory of Metal Matrix Composites, School of Chemistry & Chemical Engineering, Shanghai Jiao Tong University, 800 Dongchuan Road, Shanghai 200240, P. R. China E-mail: [email protected] Fax: +86-21-54741297
Abbreviations: FCCS, fluorescence crosscorrelation spectroscopy; FCS, fluorescence correlation spectroscopy; PIV, particle image velocimetry; QD, quantum dot; R6G, Rhodamine 6G; SPIM, single-plane illumination; TIR-FCS, total internal reflection FCS C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
FCS was first introduced to study diffusion and chemical dynamics of DNA-drug intercalation by Magde, Elson, and Webb in 1970s [4]. However, only when the optical confocal configuration was adopted in the FCS system by Rigler et al. in 1993 [5], FCS has become an advanced singlemolecule detection technique due to advantages such as high sensitivity, high spatial and temporal resolution, extremely small sample requirement, short measuring times, etc. In the confocal configuration, the laser beam is strongly focused by a high numerical aperture objective (ideally numerical aperture ⬎0.9) into a diffraction limited spot. So, its detection volume is generally less than 1.0 femtoliter that can greatly suppress the background and improve signal-to-background ratio while a micrometer pinhole, introduced in the image plane, block all fluorescence out of the focal region. Now FCS has recently experienced growing popularity in the physical, chemical, and life science, such as study on the molecular interaction, chemical kinetics, conformation dynamics, diffusion dynamics, concentration, and density of molecules in living cell [1–3, 6–13]. Since a micro total analysis system (TAS) was presented by Manz, Harrison, and Widmer et al. [14–16], a great effort has been devoted to the miniaturization of instruments. Up to now, the technology related with microchannel has been successfully applied for various chemical species including drugs, environmental pollutants, small biomolecules, and biopolymers such as DNA fragments and proteins [17, 18].
Colour Online: See the article online to view Figs. 1, 3–5, 8, 9 and 11 in colour.
www.electrophoresis-journal.com
2268
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
Figure 1. (A) A typical diagram of a single-beam FCS setup [21]. (B) Scheme of FCS curves formed by correlating the fluctuation of fluorescent intensity with lag time when fluorescent molecules diffuse through the focal volume.
The advantages of FCS can be viewed under three aspects: (i) It becomes a single-molecule detection technique due to remarkable decrease of detection volume to the point that the fluctuation of single-molecules motion in the volume is measured when the concentration deceases to nanomolar. (ii) It is an in situ, nondestructive, and minor perturbing technique on the measured systems. (iii) It provides plenty of physical or chemical information. As a result, the implementation of coupling FCS to microchannel platforms (such as CE or microfluidic electrophoresis) offers the chances to assay the complex systems, such as DNA analysis and drug screening [19, 20]. This review is organized as follows: first, the theory of FCS and its coupling with capillary or microchannel are introduced. Second, a summary is presented on some key applications based on the combination of FCS including the flow profiles measurement, confined diffusion behaviors in the microchannel, multicomponent analyses, DNA analysis, characterization of quantum dots (QDs) and viruses.
2 Theory of FCS In Fig. 1A, it is presented a typical FCS setup [21]. As the 488 nm laser beam, from an argon ion laser, is reflected into a water-immersed objective by a dichroic mirror, a very small focal volume (less than 1 femtoliter) form above the objectives. Due to the diffusion, the fluctuations in fluorescence (␦F(t)) that arise around the average fluorescence F is recorded in real time by a single-photon counting module to be correlated with a digital correlator card (Fig. 1B). In principle, the normalized autocorrelation function G( ) is defined as [4, 5]: G( ) =
␦F (t)␦F (t + ) , F (t)2
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
(1)
where ␦F(t + ) fluorescence intensity fluctuations at a time t + . The signal is analyzed with respect to its self-similarity after the lag time . The autocorrelation amplitude G(0) is therefore merely the normalized variance of the fluctuating fluorescence signal ␦F(t). When fluorescent molecules diffuse freely in the tiny detection volume with a 3D Gaussian profile, Eq. (1) would be expressed as indicated in Eq. (2): Tt e − /triplet 1 1 · · 1+ · G( ) = N 1 − Tt 1+ D 1+
1 0 z0
2
· D
.
(2)
Here, Tt is the fraction of fluorophors in the triplet state and triplet is the corresponding triplet state relaxation time. N is the average number of fluorescent molecules in the detection volume, 0 and z0 are lateral and axial radii of the detection volume, and D is the characteristic diffusion time of fluorescent molecules, which is related to the diffusion constant, D and lateral radius as shown in Eq. (3): D =
02 . 4D
(3)
In fluorescence cross-correlation spectroscopy (FCCS) technique, the fluorescence fluctuations is measured by two detection channels (denoted by g and r, respectively) [22]. The deviations from the mean in the measured fluorescence from the two channels are correlated with the normalized cross-correlation function: ␦Fg (t)␦Fr (t + ) , (4) G( ) = Fg (t)Fr (t) where Fg (t), Fr (t) are the fluorescence signals from detection channel g and r and .. stands for time averaging. ␦Fg (t) is www.electrophoresis-journal.com
Microfluidics and Miniaturization
Electrophoresis 2014, 35, 2267–2278
2269
defined as the difference of the instantaneous fluorescence from the temporal mean in channel g, ␦Fr (t + ) is defined as the difference of the instantaneous fluorescence from the temporal mean in channel-r after lag time, . When no crosstalk was present between two channels, the function is computed as shown in Eq. (5): G gr ( ) = G gr (0) ·
1 1+
D,gr
· 1+
1 0 z0
2
,
(5)
D,gr
where D,gr is the characteristic diffusion time of two dyelabeled fluorescent complex in the confocal volume. When there is active transport in the form of laminar flow such as flow in the channel, the autocorrelation function G( ) in Eq. (2) is modified by adding a flow term: Tt e − /triplet 1 1 · 1+ · G( ) = N 1 − Tt 1+ D ⎛ ⎞⎞ ⎛ 2 1 ⎜ 1 ⎟⎟ ⎜ · exp ⎝− · ⎝ ⎠⎠ , 2 f 1+ 0 D 1+ · z0 D (6) where f is the average flow time of fluorophors in uniform flow traversing through the open volume element. From this autocorrelation function, the flow time is deduced by fitting the experimental data with Eq. (6). The flow velocity Vf can be calculated from Eq. (7). [23]: V f = 0 / f .
(7)
Equation (6) is deduced based on the assumption that the flow in the microchannel is uniform and single directional in a plane perpendicular to the optical axis. The continuous hydrodynamic flow can be driven by pressure by pump or elevating reservoir, and electric field. In order to determine the flow direction in the microchannel, two-beam FCCS was combined with the microchannel. Brinkmeier et al. derived an analytical expression for the cross-correlation function taking into account that the effects of diffusion and flow as seen in Eq. (8) [6]: G( ) =
1 1 1 · · · 2 N 1+ 0 D 1+ z0 D 2 R2 2 + 1 − 2 2 cos ␣ , exp − 2 0 2f f
(8)
where N is the average number of molecules in the detection volume, and f is the average duration of the transit between the two detection volumes. The cross-correlation function reaches a maximum when is equal to f . If the direction is changed, the position and height of the maximum of the peak will be changed according to Eq. (8) (with ␣ = 0o and 180o , respectively). C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Figure 2. Schematic diagram of the CE/FCS experimental apparatus. Reprinted with permission from [24].
3 Coupling of FCS with capillary and microchannel 3.1 Capillary analytical system CE is a microscale analytical technique characterized by short analysis time, small sample and reagent requirements, and high separation efficiency. The first attempt about combination of FCS with capillary was implemented by Van Orden and Keller in 1998 [24]. An example of such system is shown in Fig. 2. It consists of an Ar ion laser beam (514 nm) passing through an oil-immersed microscope objective and focused into a window on a fused silica capillary (40 m internal diameter). Laminar flow of the analyte solution in the capillary is established by a pneumatic pressure regulator or electric fields applied between the inlet and outlet of the capillary. Different to orthogonal channel of microfluidic chip, the curvature of the capillary walls introduced distortions to the shape of the detection region from an ideal Gaussian profile due to spherical aberrations and beam astigmatism. Later, Van Orden et al. proposed the combination of two laser beams, each one focused into one window in the capillary, as shown in Fig. 3. Different from the single-beam FCS, in this new approach, denominated FCCS, the laser beam is split and then recombined into two nearly parallel beams by two 50/50 beam splitters. The resulting two quasi-parallel laser beams are reflected into the back aperture of the microscope objective by a dichroic mirror. Two nearly identical focal volumes form above the coverslip. The distance between two focal volumes is about 5 m. The excited fluorescence signal from the sample was collected by the same objective and passed through a dichroic mirror. The produced fluorescence fluctuations were cross-correlated with a two channel correlator card. www.electrophoresis-journal.com
2270
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
the observation volume created by an about 350 nm channel is 100 times smaller than observation volume using conventional FCS [30].
3.3 Other configurations for FCS with a microchannel
Figure 3. Schematic representation of coupling of two-beam FCCS into the squared capillary. Reprinted with permission from [26].
With similar method, other FCS-based techniques were also combined into the microchannel including two-focus FCCS [31–34], scanning FCS [35], total internal reflection FCS (TIRFCS or TIR-FCCS) [36–38], zero-mode waveguides FCS [39], and twin-focus photothermal correlation spectroscopy (TwinPhoCS) [40]. These FCS techniques demonstrate the versatility of characterizing on flow properties in the channel. For example, using two-focus FCCS, not only the flow velocity in the channel can be measured but also the flow direction can be discriminated. However, TIR-FCS or TIR-FCCS show unique advantage over other methods about the flow profiling in the close proximity of an interface that cannot be determined by single-beam FCS due to the resolution limit of optical confocal microscopies. In the measurement sensitivity, Twin-PhoCS showed the highest sensitivity of measurement on flow velocities in the channel among the FCS techniques. It was reported that using 14 nm sized gold nanoparticles (AuNPs) as photothermal probe the smallest velocity of 10 nm/ms can be detected [40].
4 Key applications 4.1 Confined diffusion of molecules in the narrow channel
Figure 4. Schematic setup of FCS with microchip electrophoresis. Reprinted with permission from [28].
3.2 Microchannel analytical system The first attempt about combination of FCS with a microfluidic channel was implemented by G¨osch et al. in 2000 [27]. Figure 4 is a typical scheme for the coupling of FCS and a microchip electrophoresis system [28]. Due to vertical side walls and planar bottom surface in the channel of microfluidic chip, the diffraction limited laser focus can be coupled into the channel and the position of the detection region in the channel can be controlled by positioning the objective to the channel. The PDMS/glass microchip used was fabricated by using photolithography and replicating techniques [29]. One advantage of coupling of narrow channel with FCS is that it can remarkably reduce the effective observation volume and improve the S/Ns. Craighead and co-workers found that C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Molecules diffuse freely in an unrestrained system such as the open volume in the coverslip. However, the possible boundary effects from narrow channel on diffusion behaviors of molecules become apparent and have to be taken into accounts. The influence can be investigated when focusing FCS into the narrow channel [30, 41, 42]. Foquet et al. investigated the confinement effect by determining the mobility of Alexa Fluor 488–5-dUTP dye in different width channel by FCS [30]. As shown in Fig. 5, due to the additional lateral confinement, the measured FCS curves in narrow channels were found to be deviated from the 2D standard model of autocorrelation functions. And the effective observation volume in about 350 nm width nanochannel is remarkably reduced (100fold) and the S/N increased. Petr´asek et al. measured fluorescence autocorrelation curves of 10 nM Alexa 546 dye aqueous solution in silicon oxide channels with different widths (10 and 0.6 m) [43]. It was found that the curve from the 10-m wide channel can be described by unrestricted 2D standard diffusion model. But the correlation curve from the 0.6-m wide channel exhibits significantly slower decay, which is much better described by a 1D diffusion model. And the characteristic diffusion time www.electrophoresis-journal.com
Electrophoresis 2014, 35, 2267–2278
Microfluidics and Miniaturization
2271
Figure 5. (A) Scheme of detecting single molecules flowing in the nanochannel. (B) FCS analysis of Alexa Fluor 4885-dUTP inside the narrow channel region. Reprinted with permission from [30].
of Alexa 546 in narrow channel was larger than that in the wide channel. Sanguigno et al. developed reflection and superposition method to account for the confinement effect on the diffusion behavior of Rhodamine 6G (R6G) dye in the channel with different height and width [44]. It was found that the presence of walls in the channels has a severe effect on the FCS curve shape, which results in its strong deviation from the standard FCS model. Besides, due to conformational rearrangements related to molecule flexibility and surface interactions dynamics, macromolecules presented different diffusive behavior compared to the ones of small molecules in the nanochannel. De Santo et al. investigated the influence of confinement on motion of fluorescent PEG and Rhodamine green dye in glass nanochannels (height is 10–30 nm and width is in the range of 5–30 m) with FCS [45]. It was observed that Rhodamine green appeared to be almost unaffected from confinement but all macromolecules showed a reduction of their diffusion coefficient of almost one order of magnitude.
4.2 Measurement on the flow velocity and profile in the channel 4.2.1 Flow velocity and profile in the channel The combination of FCS with a channel was originally developed to measure the flow velocity and profile in the channel [27]. They have become the important tools for flow mapping in the channel instead of conventional methods. The ability to quickly measure flow parameters in microfluidic devices is critical for TAS applications. Early, a number of methods have evolved to map fluid flow profiles in microfluidic structure such as particle image velocimetry (PIV), optical Doppler tomography, and fluorescence bleaching techniques [46–48]. In the PIV measurement, the micrometer-sized fluorescent beads are first introduced into the microstructure, and then the positions of flowing particles are recorded according to the obtained full images with a known time delay between
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
frames. The displacement of individual particles during the time delay is measured and so the velocity for a given fluid position is calculated based on the displacement and delay time. The main drawbacks in these methods are that these introduced beads can clog the microstructure. And the big size bead can disturb the fluid flow properties or lower the detection resolution. Compared with above methods, FCS has high spatial resolution requiring very short measurement times. The most important is that no beads are needed in FCS. Instead, the flow properties including the flow velocity and profiles in the channel can be measured by FCS and imaged after highly diluted concentration of fluorescent probes were introduced into the microchannel, as shown in Fig. 6 [27, 49, 50]. Application of FCS for flow measurements was first presented by Magde et al. in 1978 when R6G dye was used as probe to measure velocity in water [23]. With the development of FCS technique, G¨osch et al. firstly proposed to apply FCS to monitor the flow in the microchannel. FCS was used to scan the flow in a 50 × 50 m2 microchannel with 1 m steps in both the vertical and the horizontal directions with results showing that the flow profile in the microchannel was parabolic in both dimensions (Fig. 7) [27]. Later, Weston and co-workers used similar method to map fluid velocities and profiles of flow in PDMS/glass microfluidic devices [51, 52]. The measured fluid velocities appeared the high linearity with applied pressure over a range of velocities spanning four orders of magnitude from about 10 to more than 1200 mm/s. On the other hand, it was found that the shape of these profiles depended on the geometry of the channels. The precision mapping of flow velocity and profile in the capillary is also useful to characterize and optimize flow characteristics. Visser and co-workers used FCS to detect flowing fluorescent particles and molecules in a capillary and study optical trap forces produced by laser beam [53]. These reports indicated that the uniform fluid flow in the microchannel was driven by the mechanical force such as by elevating reservoir of diluted probe, etc. [27]. The flow properties in the channel were also investigated when the flow in the channel was driven by the imposed high electrical
www.electrophoresis-journal.com
2272
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
Figure 6. Demonstration of the ability of FCS to measure fluid flow rates over a very wide dynamic range. Panel (A) shows selected autocorrelation functions calculated from fluorescence intensity trajectories along with fits to the flow model and panel (B) shows the flow rate measured using FCS versus applied pressure. Reprinted with permission from [51].
Figure 8. (A, C) Intensity and (B, D) velocity images of a constant radius 90o turn and a 90o turn designed for reduced dispersion. Reprinted with permission from [52].
Figure 7. Flow profiles in the microchannel scanned in (A) vertical and (B) horizontal directions at an applied reservoir height of 20 cm (V = 15 mm/s). Reprinted with permission from [27].
field [54]. Rigneault et al. verified using FCS method that a uniform fluid flow was also formed in the electrophoresis channel. The flow velocity in the channel is constant for flow and the velocity increases linearly with the applied electric field. These early investigations on fluidic flow mapping are exclusively focused on straight channel in the microfluidic chip. However, besides the straight channel for sample separation, the microfluidic chip is also composed of some reservoirs, chambers, holes, and mixers connected with junction channel for other versatile applications. Due to the irregularity of C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
the structures, the velocity distribution and flow profile in the irregular microstructure are of course remarkably different to that of straight channels being more difficult to characterize. The flow profile in these cases can also be evaluated by FCS techniques. Pollack and co-workers measured the 3D flow profiles in a five-inlet port mixer with FCS when the flow was directed to the channel from five inlets at different flow rates [55]. Pappas and co-workers applied FCS to measure the flow in a microfluidic vortex formed in T-shaped intersection. The flow at different points in the T-shaped intersection was mapped. It was found that the center of the vortex presented lower flow rates than the main channel [56]. As a typical example, the velocity profiles and images of the flow in the 90o turn of microchannels are obtained with FCS as shown in Fig. 8 [52]. Besides, Carlotto et al. investigated the hydrodynamic focusing effect in an asymmetric focused flow formed in a microfluidic device by employing FCS to measure the flow speeds across the channels and the concentration profile of www.electrophoresis-journal.com
Electrophoresis 2014, 35, 2267–2278
the central focused asymmetric flux along the flow direction [57]. And it was found that experimental data agreed with the prediction of numerical simulations based on a standard Navier–Stokes finite-element approach. With the decreasing of the dimensions of the channel, FCS shows more advantages over other characterization method such as PIV. FCS can be used to measure the flow velocities in the submicrochannel and the nanochannnel. Craighead and co-workers fabricated submicrofluidic channels with two lateral dimensions smaller than 1 m in fused silica [30]. It was found that the effective dimensionality of diffusion in the channel was highly depended on the channel geometry. In the narrow channel, a 100-fold reduction in the effective volume was observed due to the additional lateral confinement. Although the size of channel decreased to several hundred nanometers, it was demonstrated that the flow speed in the nanochannel was still linearly proportional to the applied voltage [42].
4.2.2 Flow direction The classical hydrodynamic flow profile within microchannel and even in the nanochannel were accessed with single laser beam FCS or single-focus FCS. These methods have some inherent disadvantages that cannot be resolved: (i) The flow directions cannot be monitored due to its symmetric detection volume; (ii) When applied to flow profiling in the close proximity of an interface, they suffer from the limited resolution of optical microscopes. Despite that several FCS techniques can be used to monitor the flow directions. One technique is the application of line-scan FCS. Wohland and co-workers performed line-scan FCS experiments on a modified laser-scanning confocal microscope and demonstrated its applications in the analysis of flow profiles in the microchannel. Most important, the method showed the possibility to monitor the flow in small blood vessels such as living zebrafish tissue [35]. Hashmi et al. adopted line illumination and camera-based detection reminiscent of line-scanning confocal microscopy into flow measurement of FCS in the microchannel. The multiphase flows across the entire width of the microchannel were visualized with a high-speed camera and quantified with FCS. It was demonstrated that the flow velocity of particles in the microchannel at a speed up to the order of 1 cm/s can be resolved with the technique [58]. Another approach to measure the flow directions proposed by Brinkmeier et al. was to employ two-beam FCCS instead of classical single-beam FCS into the channel [6, 59]. With two-beam FCCS, fluorescent molecules successively pass the two spatially separated confocal volumes and then the signals from two volumes are cross-correlated. The maximum in a cross-correlation curve corresponds the time that it takes for fluorophore to transport from the first to the second volume. Thus, the migration velocity and direction of fluorescent molecules between two focal volumes can be measured [6, 34, 59, 60]. C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Microfluidics and Miniaturization
2273
The dual-focus FCS (2fFCS) was developed by Enderlein and co-workers [31], and it was introduced to map the 3D flow in the microchannel [33]. In contrast to two-beam FCCS, 2fFCS makes use of the intentional overlap of the two detection regions. Several experiments demonstrated its significant advantages (freedom from calibration and increased accuracy) in the measurement of flow velocities over conventional single-focus FCS [33, 34]. An important application of flow velocity measurement based on these two-beam FCCS or based on two-focus FCCS is the measurement of fast reaction kinetic parameters. Dittrich et al. developed a spatial two-beam FCCS method to accurately determine the reaction time by measuring the exact flow time from the mixing point to the observation point in a miniaturized continuous-flow (CF) reactor. Using this CF device, the kinetics of the irreversible cleavage of a DNA strain oligomer by the enzyme exonuclease was studied [61]. It is very important to converse spatial information into the respective temporal information based on the measured flow velocity by FCCS method in the investigation on the reaction kinetics. The similar approach of “position-to-time conversion” based on FCS method was also reported by Wunderlich et al. [62]. Using this approach, single-molecule kinetics of B-domain of protein A in the microfluidic mixer devices was recorded on timescales from milliseconds to minutes.
4.2.3 Flow profiling in the interface In the proximity of an interface, the normal resolution can be significantly increased using TIR microscopy. In order to overcome the disadvantage of one-beam FCS, two-beam FCCS, or line-scan FCS on flow profiling in the close proximity of an interface caused by the limited resolution of these techniques, new TIR-FCS or TIR-FCCS configurations were developed to measure velocity profiles in the interface [36–38]. For example, Yordanov et al. tuned the evanescent wave penetration depth of TIR and the velocity at different distances less than 200 nm from the interface was measured [36]. It should be pointed out that the evanescent wave illumination is one of plane illumination methods. So by adopting the TIR illumination into FCS, more comprehensive information about the flow profile can be obtained. Another illumination configuration similar to TIR-FCS can be achieved by adopting single-plane illumination (SPIM) into FCS for measurement on the flow. Wohland et al. applied SPIM as excitation source and an electron multiplying CCD (EMCCD) as array detectors, as shown in Fig. 9 [63]. Using the technique, blood flow profiles and flow directions within living zebrafish can be determined. The most importance is that the measurement depth can be adjusted in SPIM-FCCS.
4.3 Multicomponent analyses in the channel FCS is a highly sensitive and rapid single-molecule detection technique that is often used to perform multicomponent www.electrophoresis-journal.com
2274
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
Figure 9. SPIM-FCS measurements within a live zebrafish 48 h postfertilization. (A) The ACFs of 0.2 m microspheres within a blood vessel. (B) This figure shows cross-correlation functions between the central pixel of the 20 × 20 pixel region of interest and the surrounding pixels at a distance of three pixels (same experiments as A). Reprinted with permission from [63].
analysis in static solutions based on the different diffusion times of the species through the tightly focused detection volume, which scales with the cubic root of the molecular size. In order to differentiate species based on their diffusion time, a relative difference in diffusion time between two different species of at least a factor of two is required [22]. However, in many multicomponent analyses, molecular size of the analyte species is comparable. So, it is necessary to develop some methods to enhance their difference of characteristic diffusion time in focal volume. A electrophoretic mobility shift assay method by combining FCS with CE (FCS-CE) developed by Van Orden and Keller can resolve the problem above [24]. In the method, transport of mixture of R6G and R6G-labeled dCTP (R6GdCTP) nucleotides in the capillary is measured as model by FCS when they continuously flow through the capillary in the presence of an electric field. The difference of transit times between observed R6G and R6G-dCTP in the volume increase due to their different electrical mobility in the capillary. The increased difference of transit times can agree with the accuracy requirement for two component fitting procedure in FCS measurement. The multicomponent analysis procedure based on FCS-CE is accomplished to measure the relative concentration of R6G and R6G-dCTP mixture although their molecular weight difference is minor. This method combines the selectivity of CE with the sensitivity and rapid analysis times of FCS. Compared with conventional CE method, less samples volume and short analysis time are required. The assay method was extended to investigate the interaction between DNA-protein [64, 65]. LeCaptain et al. studied the interaction of single-stranded DNA-binding protein (SSB) with fluorescently labeled 39-mer single-stranded DNA (ssDNA) [64]. It was verified that the determined binding ratios agreed with the results obtained by CE separation. The resolution of the bound and unbound complexes was significantly enhanced compared to conventional FCS. Yeh et al. combined the technique with microfluidic chip electrophoresis and study the dissociation of transcription factor Sp1DNA complex by doxorubicin [65]. The binding fractions of Sp1-DNA complex at different Sp1 concentration was measured with two-component fitting procedure of FCS. The binding constant measured by FCS was found to be comparable to that measured by conventional electrophoresis assay.
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
In comparison to FCS-CE, the coupling of twobeam FCCS with microchannel electrophoresis (two-beam FCCS/CE) shows more advantage on the multicomponent analysis. For example, in the two-beam FCCS/CE, the obtained cross-correlation function contains separate peaks corresponding to the electrophoretic flow velocities of ssDNA and conjugates of SSB with ssDNA. So the relative concentrations of the bound and unbound analytes could be determined from the curve with two component analysis [25]. Multicomponent analysis procedure based on two-beam FCCS/CE has been applied for simultaneous measurement of positive and negative ions [66] and assay of protein digests [67]. The transport of positive (cationic R6G) and negative ions (anionic 5-carboxytetramethylrhodamine) in the electrophoresis capillary was monitored by two-beam FCCS/CE [66]. The obtained cross-correlation multicomponent analysis results demonstrate that the determined relative concentrations of each analyte agreed with the expected values. Brister and Weston employed two-beam FCCS to characterize the migration rates of fluorescently labeled protein digests in a microfluidic chip electrophoresis system [67]. Meanwhile, Monte Carlo simulation methods were presented to understand the capabilities and limitations of the two-beam FCCS/CE method. The experimental and simulation results demonstrated that the application of two-beam FCCS in the electrophoresis assay can aid proteomics research significantly. In these two beam-FCCS measurements, crosscorrelation curves are obtained when fluorescent molecules successively pass the two focal volumes. However, many of the detected fluorescent molecules by the first focal volume diffuse away at the second focal volume sites and hereby cannot be measured because the focal volumes are far less than the width of channel. So only part of molecules that transit both detection regions contribute to the cross-correlation peak of two beam-FCCS, which is adverse for the accumulation of signal and detection for target molecules. In order to overcome the disadvantage, a two-beam line FCCS configure was developed by Schiro et al. for use in continuous-flow single-molecule CE, where two rectangular illumination volumes completely match with the width of the microchannel, as shown in Fig. 10 [68]. The combination of this configure with CE was used to determine the mixture of FITC, FITC-labeled glycine, and FITC-labeled glutamate. In the
www.electrophoresis-journal.com
Microfluidics and Miniaturization
Electrophoresis 2014, 35, 2267–2278
Figure 10. (A) Schematic illustrating the experimental setup for continuous-flow single-molecule CE. (B) Simulation showing the distribution of laser intensity across the width of a 2-m channel for two spatially separated focal volumes. Reprinted with permission from [68].
geometry, it was observed that nearly every molecule in a sample was detected. This high detection efficiency is critical for the chemical analysis on low copies of molecules in single cells or subcellular compartments.
4.4 DNA analysis Combination of FCS with microchannel electrophoresis shows great potential in the study of fundamental physical properties of DNA undergoing electrophoretic migration [69, 70]. For example, it was observed that the electrophoretic behavior of DNA in the microchannel was highly dependent upon the electric field as the migration speed linearly increases with the applied electric field [69]. When FCCS was used to study electrophoretic behavior of DNA in the microchannel, both the translational diffusion and electrophoretic flow properties could be obtained from the measurement. For example, electrophoretic mobilities of different length of double-stranded DNA (75–1019 bp) in microfluidic channels were determined using dfFCS. It was found that when dsDNA were separated in the free solution, their mobility is independent of DNA length. However, when polyethylene oxide was sieved as the separation media, the electrophoretic rates become size dependent with a power-law exponent between 0.28 and 0.31 [70]. The role of counterions in modulating the effective charge and diffusion properties of ssDNA was investigated by C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
2275
Fogarty et al. [71]. It was observed that with the concentration increase of Mg2+ , the migration velocity of ssDNA decreased but the rate of ssDNA translational diffusion increased. The results that larger diffusion constants and smaller hydrodynamic radii indicated that the existence of Mg2+ neutralized the negative charges on the ssDNA backbone, which lowers the self-repulsion between nucleotides and makes the ssDNA form more compact structures. The kinetics and energetics of DNA hairpin folding was investigated by Jung et al. using simultaneous two-beam FCCS and single-beam FCS [26,72]. In the hairpin DNA, each end is attached with a dye and a quencher. The folding and unfolding of hairpin DNA results in intensity fluctuation due to the fluorescence on/off characteristic caused by the contact between the fluorophor and the quencher. The correlation function determined with FCS thus contains fluctuation contributions characterized by a diffusional relaxation time, d , and a relaxation time for conformational fluctuations, R . But it is difficult for conventional FCS to resolve two relaxation times because the relaxation process occurs on a time scale similar to or less than the sampling interval. Jung et al. successfully decoupled the different relaxation times of hairpin DNA by analyzing fluorescence fluctuations of hairpin DNA in a capillary observed at two spatially offset detection volumes with two-beam FCCS. The results showed a three-state mechanism was involved in the DNA hairpin folding reaction and a long-lived intermediate form of the DNA hairpin was included in the reaction.
4.5 Characterization on some parameters of nanoparticles Ren and co-workers developed a method by combining FCS with microfluidic chip electrophoresis to measure the surface charge of aqueous mercaptopropionic acid (MPA)-coated cadmium telluride (CdTe) QDs as shown in Fig. 3 [28]. According to the Stokes–Einstein equation, diffusion coefficients of spherical particles, D are inversely proportional to its hydrodynamic radius, R as indicated in Eq. (9): D=
kT , 6R
(9)
where kT is the thermal energy and is the viscosity of the solution. Thus, according to Eqs. (3) and (9), the hydrodynamic radius, R of QDs can be calculated as indicated in Eq. (10): R=
2kT D . 302
(10)
As shown in Eq. (7), migration velocity, Vf of QDs in the channel can be calculated based on the parameters (0 and f ) by fitting FCS raw data with Eq. (6). QDs migrates in the microfluidic channel with no electrosmotic flow under the electric field, so their migration velocity Vf is positively www.electrophoresis-journal.com
2276
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
tures. It is expected that the approach can be straightforwardly extended to simultaneous detection of several virus types.
5 Conclusion
Figure 11. Normalized FCS curves of aqueous CdTe QDs in the microchannel under the different electric field strength. Reprinted with permission from [28].
proportional to the intensity of electric field E and can be expressed as indicated in Eq. (11): q E, (11) Vf = 6R where q is the surface charge of fluorescent particles, and E the intensity of electric field. So, when R in Eq. (11) is replaced with one in Eq. (10) and Eq. (11) is combined with Eq. (7), the relationship between the average migration time f and the intensity of electric field E can be deduced as Eq. (12): 0 q 1 = E = s × E. f 4kT D
(12)
As a result, the charge q can be calculated on the slope (s) from the linear curve of 1/ f versus E (q = 4ksT D /0 ). Figure 11 is the normalized fluorescence autocorrelation function of QDs under the different electric field strength. It demonstrated that the transit time decreased with the increased electric field strength. The measured results show that the charges of QDs increased with pH value of solution. When surface ligand-mercaptopropionic acid was replaced by glutathione, the surface charge of CdTe QDs was increased by about 55.3%. This result showed that the surface charge of QDs was remarkably associated with the type of stabilizers on QDs surface, buffer pH, and other factors.
The applications of combination of single-molecule detection technique—FCS with channel showed more and more advantages over the sole channel separation methods. (i) Lower sample consumption. The technique of combination of FCS with channel assay has extremely high sensitivity. Single molecules are detected when they are separated in the channel. Herein, it can lower the requirement for sample volume and concentration. It is critical for the biochemical assay of low copies of molecules in single cells. (ii) More temporal information. Compared with the conventional channel separation method such as microfluidic chip electrophoresis, implementation of FCS in channel can provide the dynamics information in the time scale from microsecond to second such as the microsecond relaxation and translation of molecules. (iii) Enhanced spatial information. Narrow channels are more regarded as a miniaturized reaction and separation platform. The combination of FCS can provide the 3D spatial information about reaction and separation in the channels. It can be concluded that the combination strategies is emerging but important in the analytical chemistry, analytical biochemistry, high-throughput drug screening, and in vivo study of living cells. This work was financially supported by NFSC (20905048, 21075081, 21135004, and 21327004), Innovation Program of Shanghai Municipal Education Commission (14ZZ024), Shanghai Educational Development Foundation (2008CG12), and SMC-Chenxin Young Scholar project sponsored by Shanghai Jiao Tong University. The authors have declared no conflict of interest.
6 References [1] Punj, D., Mivelle, M., Moparthi, S. B., Van Zanten, T. S., Rigneault, H., Van Hulst, N. F., Garcia-Parajo, M. F., Wenger, J., Nat. Nanotechnol. 2013, 8, 512–516. [2] Kim, S. A., Heinze, K. G., Schwille, P., Nat. Methods 2007, 4, 963–973. [3] Zamir, E., Lommerse, P. H. M., Kinkhabwala, A., Grecco, H. E., Bastiaens, P. I. H., Nat. Methods 2010, 7, 295–298.
4.6 Detection of viruses Rudenko et al. demonstrate the possibility to detect the Q bacteriophage on the single-virus level by integrating microfluidic chip with FCS system [73]. Multiparameters on viruses including diffusion coefficient, flow velocity, and concentration of viruses were extracted from FCS experiments. Using the approach, the signal of single phage can be distinguished from their structural parts, agglomerates, or mix C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
[4] Magde, D., Elson, E. L., Webb, W. W., Phys. Rev. Lett. 1972, 29, 705–708. ¨ Widengren, J., Kask, P., Eur. Biophys. [5] Rigler, R., Mets, U., J. 1993, 22, 169–175. [6] Brinkmeier, M., Dorre, K., Stephan, J., Eigen, M., Anal. Chem. 1999, 71, 609–616. [7] Rigler, R., Biochem. Biophys. Res. Commun. 2010, 396, 170–175. [8] Elson, E. L., Biophys. J. 2011, 101, 2855–2870.
www.electrophoresis-journal.com
Electrophoresis 2014, 35, 2267–2278
Microfluidics and Miniaturization
2277
[9] Ruan, L. G., Xu, Z. C., Lan, T., Wang, J. J., Liu, H., Li, C. D., Dong, C. Q., Ren, J. C., Anal. Chem. 2012, 84, 7350–7358.
[36] Yordanov, S., Best, A., Butt, H. J., Koynov, K., Opt. Express 2009, 17, 21149–21158.
[10] Dong, C. Q., Chowdhury, B., Irudayaraj, J., Analyst 2013, 138, 2871–2876.
[37] Blom, H., Hassler, K., Chmyrov, A., Widengren, J., Int. J. Mol. Sci. 2010, 11, 386–406.
[11] Dong, C. Q., Qian, H. F., Fang, N. H., Ren, J. C., J. Phys. Chem. B 2006, 110, 11069–11075.
[38] Schmitz, R., Yordanov, S., Butt, H. J., Koynov, K., Dunweg, B., Phys. Rev. E 2011, 84, 066306.
[12] Dong, C. Q., Qian, H. F., Fang, N. H., Ren, J. C., J. Phys. Chem. C 2007, 111, 7918–7923.
[39] Liao, D., Galajda, P., Riehn, R., Ilic, R., Puchalla, J. L., Yu, H. G., Craighead, H. G., Austin, R. H., Opt. Express 2008, 16, 10077–10090.
[13] Tang, L. C., Dong, C. Q., Ren, J. C., Talanta 2010, 81, 1560–1567. [14] Manz, A., Graber, N., Widmer, H. M., Sens. Actuators B. Chem. 1990, 1, 244–248. [15] Harrison, D. J., Manz, A., Fan, Z., Luedi, H., Widmer, H. M., Anal. Chem. 1992, 64, 1926–1932.
[40] Selmke, M., Schachoff, R., Braun, M., Cichos, F., RSC Adv. 2013, 3, 394–400. [41] Gennerich, A., Schild, D., Biophys. J. 2000, 79, 3294–3306.
[16] Harrison, D. J., Fluri, K., Seiler, K., Fan, Z., Effenhauser, C. S., Manz, A., Science 1993, 261, 895–897.
[42] 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.
[17] Arora, A., Simone, G., Salieb-Beugelaar, G. B., Kim, J. T., Manz, A., Anal. Chem. 2010, 82, 4830–4847.
[43] Petrasek, Z., Krishnan, M., Monch, I., Schwille, P., Microsc. Res. Techniq. 2007, 70, 459–466.
[18] Kovarik, M. L., Ornoff, D. M., Melvin, A. T., Dobes, N. C., Wang, Y., Dickinson, A. J., Gach, P. C., Shah, P. K., Allbritton, N. L., Anal. Chem. 2013, 85, 451–472.
[44] Sanguigno, L., De Santo, I., Causa, F., Netti, P., Anal. Chem. 2010, 82, 9663–9670.
[19] Dittrich, P. S., Manz, A., Nat. Rev. Drug Discov. 2006, 5, 210–218.
[46] Meinhart, C. D., Wereley, S. T., Santiago, J. G., Exp. Fluids 1999, 27, 414–419.
[20] Fogarty, K., Van Orden, A., Methods 2009, 47, 151–158. [21] Dong, C. Q., Ren, J. C., Analyst 2010, 135, 1395–1399.
[47] Devasenathipathy, S., Santiago, J. G., Takehara, K., Anal. Chem. 2002, 74, 3704–3713.
[22] Schwille, P., Meyer-Almes, F. J., Rigler, R., Biophys. J. 1997, 72, 1878–1886.
[48] Gai, H. W., Li, Y., Silber-Li, Z., Ma, Y. F., Lin, B. C., Lab Chip 2005, 5, 443–449.
[23] Magde, D., Web, W. W., Elson, E. L., Biopolymers 1978, 17, 361–367.
[49] Edel, J. B., Hill, E. K., de Mello, A. J., Analyst 2001, 126, 1953–1957.
[24] Van Orden, A., Keller, R. A., Anal. Chem. 1998, 70, 4463–4471.
[50] Liu, T. C., Wohland, T., Watt, F., Bettiol, A., Udalagama, C., Wang, L. P., Med. Phys. 2012, 39, 5060–5064.
[25] LeCaptain, D. J., Van Orden, A., Anal. Chem. 2002, 74, 1171–1176.
[51] Brister, P. C., Kuricheti, K. K., Buschmann, V., Weston, K. D., Lab Chip 2005, 5, 785–791.
[26] Jung, J., Van Orden, A., J. Am. Chem. Soc. 2006, 128, 1240–1249.
[52] Kuricheti, K. K., Buschmann, V., Weston, K. D., Appl. Spectrosc. 2004, 58, 1180–1186.
¨ [27] Gosch, M., Blom, H., Holm, J., Heino, T., Rigler, R., Anal. Chem. 2000, 72, 3260–3265.
[53] Kunst, B. H., Schots, A., Visser, A. J. W. G., Anal. Chem. 2002, 74, 5350–5357.
[28] Dong, C. Q., Bi, R., Qian, H. F., Li, L., Ren, J. C., Small 2006, 2, 534–538.
[54] Lenne, P. F., Colombo, D., Giovannini, H., Rigneault, H., Single Mol. 2002, 3, 194–200.
[29] Chen, L., Ren, J. C., Bi, R., Chen, D., Electrophoresis 2004, 25, 914–921.
[55] Park, H. Y., Qiu, X. Y., Rhoades, E., Korlach, J., Kwok, L. W., Zipfel, W. R., Webb, W. W., Pollack, L., Anal. Chem. 2006, 78, 4465–4473.
[30] Foquet, M., Korlach, J., Zipfel, W. R., Webb, W. W., Craighead, H. G., Anal. Chem. 2004, 76, 1618–1626. [31] Dertinger, T., Pacheco, V., von der Hocht, I., Hartmann, R., Gregor, I., Enderlein, J., Chemphyschem 2007, 8, 433–443. [32] Felten, M., Staroske, W., Jaeger, M. S., Schwille, P., Duschl, C., Electrophoresis 2008, 29, 2987–2996. [33] Arbour, T. J., Enderlein, J., Lab Chip 2010, 10, 1286–1292.
[45] De Santo, I., Causa, F., Netti, P. A., Anal. Chem. 2010, 82, 997–1005.
[56] Liu, K., Tian, Y., Burrows, S. M., Reif, R. D., Pappas, D., Anal. Chim. Acta 2009, 651, 85–90. [57] Carlotto, S., Fortunati, I., Ferrante, C., Schwille, P., Polimeno, A., Microfluid Nanofluid 2011, 10, 551–561. [58] Hashmi, S. M., Loewenberg, M., Dufresne, E. R., Opt. Express 2007, 15, 6528–6533. [59] Brinkmeier, M., Dorre, K., Riebeseel, K., Rigler, R., Biophys. Chem. 1997, 66, 229–239.
[34] Lumma, D., Best, A., Gansen, A., Feuillebois, F., Radler, J. O., Vinogradova, O. I., Phys. Rev. E 2003, 67, 056313.
[60] Dittrich, P. S., Schwille, P., Anal. Chem. 2002, 74, 4472–4479.
[35] Pan, X. T., Yu, H., Shi, X. K., Korzh, V., Wohland, T., J. Biomed. Opt. 2007, 12, 014034.
[61] Dittrich, P. S., Muller, B., Schwille, P., Phys. Chem. Chem. Phys. 2004, 6, 4416–4420.
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
www.electrophoresis-journal.com
2278
C. Dong and J. Ren
[62] Wunderlich, B., Nettels, D., Benke, S., Clark, J., Weidner, S., Hofmann, H., Pfeil, S. H., Schuler, B., Nat. Protoc. 2013, 8, 1459–1474. [63] Wohland, T., Shi, X. K., Sankaran, J., Stelzer, E. H. K., Opt. Express 2010, 18, 10627–10641. [64] LeCaptain, D. J., Michel, M. A., Van Orden, A., Analyst 2001, 126, 1279–1284. [65] Yeh, H. C., Puleo, C. M., Lim, T. C., Ho, Y. P., Giza, P. E., Huang, R. C., Wang, T. H., Nucleic Acids Res. 2006, 34, e144. [66] Fogarty, K., Van Orden, A., Anal. Chem. 2003, 75, 6634–6641. [67] Brister, P. C., Weston, K. D., Analyst 2006, 131, 303–310.
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Electrophoresis 2014, 35, 2267–2278
[68] Schiro, P. G., Kuyper, C. L., Chiu, D. T., Electrophoresis 2007, 28, 2430–2438. [69] Foquet, M., Korlach, J., Zipfel, W., Webb, W. W., Craighead, H. G., Anal. Chem. 2002, 74, 1415–1422. [70] Bayer, J., Radler, J. O., Electrophoresis 2006, 27, 3952–3963. [71] Fogarty, K., McPhee, J. T., Scott, E., Van Orden, A., Anal. Chem. 2009, 81, 465–472. [72] Jung, J., Ihly, R., Scott, E., Yu, M., Van Orden, A., J. Phys. Chem. B 2008, 112, 127–133. [73] Rudenko, M. I., Kuhn, S., Lunt, E. J., Deamer, D. W., Hawkins, A. R., Schmidt, H., Biosens. Bioelectron. 2009, 24, 3258–3263.
www.electrophoresis-journal.com
Electrophoresis 2014, 35, 2267–2278
Chaoqing Dong Jicun Ren State Key Laboratory of Metal Matrix Composites, School of Chemistry & Chemical Engineering, Shanghai Jiao Tong University, Shanghai, P. R. China
Received December 24, 2013 Revised March 10, 2014 Accepted March 21, 2014
Review
Coupling of fluorescence correlation spectroscopy with capillary and microchannel analytical systems and its applications In the past decade, fluorescence correlation spectroscopy (FCS) has become an ultrasensitive and noninvasive single-molecule detection technique, which is widely applied in the physical, chemical, and life-science research. The coupling of FCS with narrow channel flow systems including the ones based on capillary provide the important, convenient, and sensitive assay platforms for probing and understanding the behavior of single molecules or nanoparticles with improved temporal and spatial resolution and need for less sample volume among other advantages. This review focus on different approaches for FCS with capillary and microchannel analytical systems and its applications in confined diffusion study, flow profiles, and imaging of narrow channel, multicomponent analyses such as protein, DNA analysis, and characterization on nanoparticles. Keywords: Capillary electrophoresis / Coupling / Fluorescence correlation spectroscopy / TAS DOI 10.1002/elps.201300648
1 Introduction Fluorescence correlation spectroscopy (FCS) is an ultrasensitive and noninvasive technique for single-molecule detection that uses statistical analysis of the fluorescence fluctuation emitted from a small and optically well-defined open volume element [1–3]. In contrast to other fluorescence techniques, FCS can provide not only the emission intensity of fluorophors in the volume, but also other important parameters such as the diffusion or mobility coefficients of fluorophors, etc. because of the minor intensity fluctuations caused by the minute deviations of molecules from thermal equilibrium. Thus, any factors that lead to the deviations from equilibrium can be investigated by FCS including diffusion, physical or chemical reactions, aggregation, etc. Using FCS, local concentration, diffusion coefficients, chemical reaction rate as well as photophysical properties of fluorophors can be determined.
Correspondence: Professor Jicun Ren, State Key Laboratory of Metal Matrix Composites, School of Chemistry & Chemical Engineering, Shanghai Jiao Tong University, 800 Dongchuan Road, Shanghai 200240, P. R. China E-mail: [email protected] Fax: +86-21-54741297
Abbreviations: FCCS, fluorescence crosscorrelation spectroscopy; FCS, fluorescence correlation spectroscopy; PIV, particle image velocimetry; QD, quantum dot; R6G, Rhodamine 6G; SPIM, single-plane illumination; TIR-FCS, total internal reflection FCS C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
FCS was first introduced to study diffusion and chemical dynamics of DNA-drug intercalation by Magde, Elson, and Webb in 1970s [4]. However, only when the optical confocal configuration was adopted in the FCS system by Rigler et al. in 1993 [5], FCS has become an advanced singlemolecule detection technique due to advantages such as high sensitivity, high spatial and temporal resolution, extremely small sample requirement, short measuring times, etc. In the confocal configuration, the laser beam is strongly focused by a high numerical aperture objective (ideally numerical aperture ⬎0.9) into a diffraction limited spot. So, its detection volume is generally less than 1.0 femtoliter that can greatly suppress the background and improve signal-to-background ratio while a micrometer pinhole, introduced in the image plane, block all fluorescence out of the focal region. Now FCS has recently experienced growing popularity in the physical, chemical, and life science, such as study on the molecular interaction, chemical kinetics, conformation dynamics, diffusion dynamics, concentration, and density of molecules in living cell [1–3, 6–13]. Since a micro total analysis system (TAS) was presented by Manz, Harrison, and Widmer et al. [14–16], a great effort has been devoted to the miniaturization of instruments. Up to now, the technology related with microchannel has been successfully applied for various chemical species including drugs, environmental pollutants, small biomolecules, and biopolymers such as DNA fragments and proteins [17, 18].
Colour Online: See the article online to view Figs. 1, 3–5, 8, 9 and 11 in colour.
www.electrophoresis-journal.com
2268
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
Figure 1. (A) A typical diagram of a single-beam FCS setup [21]. (B) Scheme of FCS curves formed by correlating the fluctuation of fluorescent intensity with lag time when fluorescent molecules diffuse through the focal volume.
The advantages of FCS can be viewed under three aspects: (i) It becomes a single-molecule detection technique due to remarkable decrease of detection volume to the point that the fluctuation of single-molecules motion in the volume is measured when the concentration deceases to nanomolar. (ii) It is an in situ, nondestructive, and minor perturbing technique on the measured systems. (iii) It provides plenty of physical or chemical information. As a result, the implementation of coupling FCS to microchannel platforms (such as CE or microfluidic electrophoresis) offers the chances to assay the complex systems, such as DNA analysis and drug screening [19, 20]. This review is organized as follows: first, the theory of FCS and its coupling with capillary or microchannel are introduced. Second, a summary is presented on some key applications based on the combination of FCS including the flow profiles measurement, confined diffusion behaviors in the microchannel, multicomponent analyses, DNA analysis, characterization of quantum dots (QDs) and viruses.
2 Theory of FCS In Fig. 1A, it is presented a typical FCS setup [21]. As the 488 nm laser beam, from an argon ion laser, is reflected into a water-immersed objective by a dichroic mirror, a very small focal volume (less than 1 femtoliter) form above the objectives. Due to the diffusion, the fluctuations in fluorescence (␦F(t)) that arise around the average fluorescence F is recorded in real time by a single-photon counting module to be correlated with a digital correlator card (Fig. 1B). In principle, the normalized autocorrelation function G( ) is defined as [4, 5]: G( ) =
␦F (t)␦F (t + ) , F (t)2
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
(1)
where ␦F(t + ) fluorescence intensity fluctuations at a time t + . The signal is analyzed with respect to its self-similarity after the lag time . The autocorrelation amplitude G(0) is therefore merely the normalized variance of the fluctuating fluorescence signal ␦F(t). When fluorescent molecules diffuse freely in the tiny detection volume with a 3D Gaussian profile, Eq. (1) would be expressed as indicated in Eq. (2): Tt e − /triplet 1 1 · · 1+ · G( ) = N 1 − Tt 1+ D 1+
1 0 z0
2
· D
.
(2)
Here, Tt is the fraction of fluorophors in the triplet state and triplet is the corresponding triplet state relaxation time. N is the average number of fluorescent molecules in the detection volume, 0 and z0 are lateral and axial radii of the detection volume, and D is the characteristic diffusion time of fluorescent molecules, which is related to the diffusion constant, D and lateral radius as shown in Eq. (3): D =
02 . 4D
(3)
In fluorescence cross-correlation spectroscopy (FCCS) technique, the fluorescence fluctuations is measured by two detection channels (denoted by g and r, respectively) [22]. The deviations from the mean in the measured fluorescence from the two channels are correlated with the normalized cross-correlation function: ␦Fg (t)␦Fr (t + ) , (4) G( ) = Fg (t)Fr (t) where Fg (t), Fr (t) are the fluorescence signals from detection channel g and r and .. stands for time averaging. ␦Fg (t) is www.electrophoresis-journal.com
Microfluidics and Miniaturization
Electrophoresis 2014, 35, 2267–2278
2269
defined as the difference of the instantaneous fluorescence from the temporal mean in channel g, ␦Fr (t + ) is defined as the difference of the instantaneous fluorescence from the temporal mean in channel-r after lag time, . When no crosstalk was present between two channels, the function is computed as shown in Eq. (5): G gr ( ) = G gr (0) ·
1 1+
D,gr
· 1+
1 0 z0
2
,
(5)
D,gr
where D,gr is the characteristic diffusion time of two dyelabeled fluorescent complex in the confocal volume. When there is active transport in the form of laminar flow such as flow in the channel, the autocorrelation function G( ) in Eq. (2) is modified by adding a flow term: Tt e − /triplet 1 1 · 1+ · G( ) = N 1 − Tt 1+ D ⎛ ⎞⎞ ⎛ 2 1 ⎜ 1 ⎟⎟ ⎜ · exp ⎝− · ⎝ ⎠⎠ , 2 f 1+ 0 D 1+ · z0 D (6) where f is the average flow time of fluorophors in uniform flow traversing through the open volume element. From this autocorrelation function, the flow time is deduced by fitting the experimental data with Eq. (6). The flow velocity Vf can be calculated from Eq. (7). [23]: V f = 0 / f .
(7)
Equation (6) is deduced based on the assumption that the flow in the microchannel is uniform and single directional in a plane perpendicular to the optical axis. The continuous hydrodynamic flow can be driven by pressure by pump or elevating reservoir, and electric field. In order to determine the flow direction in the microchannel, two-beam FCCS was combined with the microchannel. Brinkmeier et al. derived an analytical expression for the cross-correlation function taking into account that the effects of diffusion and flow as seen in Eq. (8) [6]: G( ) =
1 1 1 · · · 2 N 1+ 0 D 1+ z0 D 2 R2 2 + 1 − 2 2 cos ␣ , exp − 2 0 2f f
(8)
where N is the average number of molecules in the detection volume, and f is the average duration of the transit between the two detection volumes. The cross-correlation function reaches a maximum when is equal to f . If the direction is changed, the position and height of the maximum of the peak will be changed according to Eq. (8) (with ␣ = 0o and 180o , respectively). C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Figure 2. Schematic diagram of the CE/FCS experimental apparatus. Reprinted with permission from [24].
3 Coupling of FCS with capillary and microchannel 3.1 Capillary analytical system CE is a microscale analytical technique characterized by short analysis time, small sample and reagent requirements, and high separation efficiency. The first attempt about combination of FCS with capillary was implemented by Van Orden and Keller in 1998 [24]. An example of such system is shown in Fig. 2. It consists of an Ar ion laser beam (514 nm) passing through an oil-immersed microscope objective and focused into a window on a fused silica capillary (40 m internal diameter). Laminar flow of the analyte solution in the capillary is established by a pneumatic pressure regulator or electric fields applied between the inlet and outlet of the capillary. Different to orthogonal channel of microfluidic chip, the curvature of the capillary walls introduced distortions to the shape of the detection region from an ideal Gaussian profile due to spherical aberrations and beam astigmatism. Later, Van Orden et al. proposed the combination of two laser beams, each one focused into one window in the capillary, as shown in Fig. 3. Different from the single-beam FCS, in this new approach, denominated FCCS, the laser beam is split and then recombined into two nearly parallel beams by two 50/50 beam splitters. The resulting two quasi-parallel laser beams are reflected into the back aperture of the microscope objective by a dichroic mirror. Two nearly identical focal volumes form above the coverslip. The distance between two focal volumes is about 5 m. The excited fluorescence signal from the sample was collected by the same objective and passed through a dichroic mirror. The produced fluorescence fluctuations were cross-correlated with a two channel correlator card. www.electrophoresis-journal.com
2270
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
the observation volume created by an about 350 nm channel is 100 times smaller than observation volume using conventional FCS [30].
3.3 Other configurations for FCS with a microchannel
Figure 3. Schematic representation of coupling of two-beam FCCS into the squared capillary. Reprinted with permission from [26].
With similar method, other FCS-based techniques were also combined into the microchannel including two-focus FCCS [31–34], scanning FCS [35], total internal reflection FCS (TIRFCS or TIR-FCCS) [36–38], zero-mode waveguides FCS [39], and twin-focus photothermal correlation spectroscopy (TwinPhoCS) [40]. These FCS techniques demonstrate the versatility of characterizing on flow properties in the channel. For example, using two-focus FCCS, not only the flow velocity in the channel can be measured but also the flow direction can be discriminated. However, TIR-FCS or TIR-FCCS show unique advantage over other methods about the flow profiling in the close proximity of an interface that cannot be determined by single-beam FCS due to the resolution limit of optical confocal microscopies. In the measurement sensitivity, Twin-PhoCS showed the highest sensitivity of measurement on flow velocities in the channel among the FCS techniques. It was reported that using 14 nm sized gold nanoparticles (AuNPs) as photothermal probe the smallest velocity of 10 nm/ms can be detected [40].
4 Key applications 4.1 Confined diffusion of molecules in the narrow channel
Figure 4. Schematic setup of FCS with microchip electrophoresis. Reprinted with permission from [28].
3.2 Microchannel analytical system The first attempt about combination of FCS with a microfluidic channel was implemented by G¨osch et al. in 2000 [27]. Figure 4 is a typical scheme for the coupling of FCS and a microchip electrophoresis system [28]. Due to vertical side walls and planar bottom surface in the channel of microfluidic chip, the diffraction limited laser focus can be coupled into the channel and the position of the detection region in the channel can be controlled by positioning the objective to the channel. The PDMS/glass microchip used was fabricated by using photolithography and replicating techniques [29]. One advantage of coupling of narrow channel with FCS is that it can remarkably reduce the effective observation volume and improve the S/Ns. Craighead and co-workers found that C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Molecules diffuse freely in an unrestrained system such as the open volume in the coverslip. However, the possible boundary effects from narrow channel on diffusion behaviors of molecules become apparent and have to be taken into accounts. The influence can be investigated when focusing FCS into the narrow channel [30, 41, 42]. Foquet et al. investigated the confinement effect by determining the mobility of Alexa Fluor 488–5-dUTP dye in different width channel by FCS [30]. As shown in Fig. 5, due to the additional lateral confinement, the measured FCS curves in narrow channels were found to be deviated from the 2D standard model of autocorrelation functions. And the effective observation volume in about 350 nm width nanochannel is remarkably reduced (100fold) and the S/N increased. Petr´asek et al. measured fluorescence autocorrelation curves of 10 nM Alexa 546 dye aqueous solution in silicon oxide channels with different widths (10 and 0.6 m) [43]. It was found that the curve from the 10-m wide channel can be described by unrestricted 2D standard diffusion model. But the correlation curve from the 0.6-m wide channel exhibits significantly slower decay, which is much better described by a 1D diffusion model. And the characteristic diffusion time www.electrophoresis-journal.com
Electrophoresis 2014, 35, 2267–2278
Microfluidics and Miniaturization
2271
Figure 5. (A) Scheme of detecting single molecules flowing in the nanochannel. (B) FCS analysis of Alexa Fluor 4885-dUTP inside the narrow channel region. Reprinted with permission from [30].
of Alexa 546 in narrow channel was larger than that in the wide channel. Sanguigno et al. developed reflection and superposition method to account for the confinement effect on the diffusion behavior of Rhodamine 6G (R6G) dye in the channel with different height and width [44]. It was found that the presence of walls in the channels has a severe effect on the FCS curve shape, which results in its strong deviation from the standard FCS model. Besides, due to conformational rearrangements related to molecule flexibility and surface interactions dynamics, macromolecules presented different diffusive behavior compared to the ones of small molecules in the nanochannel. De Santo et al. investigated the influence of confinement on motion of fluorescent PEG and Rhodamine green dye in glass nanochannels (height is 10–30 nm and width is in the range of 5–30 m) with FCS [45]. It was observed that Rhodamine green appeared to be almost unaffected from confinement but all macromolecules showed a reduction of their diffusion coefficient of almost one order of magnitude.
4.2 Measurement on the flow velocity and profile in the channel 4.2.1 Flow velocity and profile in the channel The combination of FCS with a channel was originally developed to measure the flow velocity and profile in the channel [27]. They have become the important tools for flow mapping in the channel instead of conventional methods. The ability to quickly measure flow parameters in microfluidic devices is critical for TAS applications. Early, a number of methods have evolved to map fluid flow profiles in microfluidic structure such as particle image velocimetry (PIV), optical Doppler tomography, and fluorescence bleaching techniques [46–48]. In the PIV measurement, the micrometer-sized fluorescent beads are first introduced into the microstructure, and then the positions of flowing particles are recorded according to the obtained full images with a known time delay between
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
frames. The displacement of individual particles during the time delay is measured and so the velocity for a given fluid position is calculated based on the displacement and delay time. The main drawbacks in these methods are that these introduced beads can clog the microstructure. And the big size bead can disturb the fluid flow properties or lower the detection resolution. Compared with above methods, FCS has high spatial resolution requiring very short measurement times. The most important is that no beads are needed in FCS. Instead, the flow properties including the flow velocity and profiles in the channel can be measured by FCS and imaged after highly diluted concentration of fluorescent probes were introduced into the microchannel, as shown in Fig. 6 [27, 49, 50]. Application of FCS for flow measurements was first presented by Magde et al. in 1978 when R6G dye was used as probe to measure velocity in water [23]. With the development of FCS technique, G¨osch et al. firstly proposed to apply FCS to monitor the flow in the microchannel. FCS was used to scan the flow in a 50 × 50 m2 microchannel with 1 m steps in both the vertical and the horizontal directions with results showing that the flow profile in the microchannel was parabolic in both dimensions (Fig. 7) [27]. Later, Weston and co-workers used similar method to map fluid velocities and profiles of flow in PDMS/glass microfluidic devices [51, 52]. The measured fluid velocities appeared the high linearity with applied pressure over a range of velocities spanning four orders of magnitude from about 10 to more than 1200 mm/s. On the other hand, it was found that the shape of these profiles depended on the geometry of the channels. The precision mapping of flow velocity and profile in the capillary is also useful to characterize and optimize flow characteristics. Visser and co-workers used FCS to detect flowing fluorescent particles and molecules in a capillary and study optical trap forces produced by laser beam [53]. These reports indicated that the uniform fluid flow in the microchannel was driven by the mechanical force such as by elevating reservoir of diluted probe, etc. [27]. The flow properties in the channel were also investigated when the flow in the channel was driven by the imposed high electrical
www.electrophoresis-journal.com
2272
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
Figure 6. Demonstration of the ability of FCS to measure fluid flow rates over a very wide dynamic range. Panel (A) shows selected autocorrelation functions calculated from fluorescence intensity trajectories along with fits to the flow model and panel (B) shows the flow rate measured using FCS versus applied pressure. Reprinted with permission from [51].
Figure 8. (A, C) Intensity and (B, D) velocity images of a constant radius 90o turn and a 90o turn designed for reduced dispersion. Reprinted with permission from [52].
Figure 7. Flow profiles in the microchannel scanned in (A) vertical and (B) horizontal directions at an applied reservoir height of 20 cm (V = 15 mm/s). Reprinted with permission from [27].
field [54]. Rigneault et al. verified using FCS method that a uniform fluid flow was also formed in the electrophoresis channel. The flow velocity in the channel is constant for flow and the velocity increases linearly with the applied electric field. These early investigations on fluidic flow mapping are exclusively focused on straight channel in the microfluidic chip. However, besides the straight channel for sample separation, the microfluidic chip is also composed of some reservoirs, chambers, holes, and mixers connected with junction channel for other versatile applications. Due to the irregularity of C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
the structures, the velocity distribution and flow profile in the irregular microstructure are of course remarkably different to that of straight channels being more difficult to characterize. The flow profile in these cases can also be evaluated by FCS techniques. Pollack and co-workers measured the 3D flow profiles in a five-inlet port mixer with FCS when the flow was directed to the channel from five inlets at different flow rates [55]. Pappas and co-workers applied FCS to measure the flow in a microfluidic vortex formed in T-shaped intersection. The flow at different points in the T-shaped intersection was mapped. It was found that the center of the vortex presented lower flow rates than the main channel [56]. As a typical example, the velocity profiles and images of the flow in the 90o turn of microchannels are obtained with FCS as shown in Fig. 8 [52]. Besides, Carlotto et al. investigated the hydrodynamic focusing effect in an asymmetric focused flow formed in a microfluidic device by employing FCS to measure the flow speeds across the channels and the concentration profile of www.electrophoresis-journal.com
Electrophoresis 2014, 35, 2267–2278
the central focused asymmetric flux along the flow direction [57]. And it was found that experimental data agreed with the prediction of numerical simulations based on a standard Navier–Stokes finite-element approach. With the decreasing of the dimensions of the channel, FCS shows more advantages over other characterization method such as PIV. FCS can be used to measure the flow velocities in the submicrochannel and the nanochannnel. Craighead and co-workers fabricated submicrofluidic channels with two lateral dimensions smaller than 1 m in fused silica [30]. It was found that the effective dimensionality of diffusion in the channel was highly depended on the channel geometry. In the narrow channel, a 100-fold reduction in the effective volume was observed due to the additional lateral confinement. Although the size of channel decreased to several hundred nanometers, it was demonstrated that the flow speed in the nanochannel was still linearly proportional to the applied voltage [42].
4.2.2 Flow direction The classical hydrodynamic flow profile within microchannel and even in the nanochannel were accessed with single laser beam FCS or single-focus FCS. These methods have some inherent disadvantages that cannot be resolved: (i) The flow directions cannot be monitored due to its symmetric detection volume; (ii) When applied to flow profiling in the close proximity of an interface, they suffer from the limited resolution of optical microscopes. Despite that several FCS techniques can be used to monitor the flow directions. One technique is the application of line-scan FCS. Wohland and co-workers performed line-scan FCS experiments on a modified laser-scanning confocal microscope and demonstrated its applications in the analysis of flow profiles in the microchannel. Most important, the method showed the possibility to monitor the flow in small blood vessels such as living zebrafish tissue [35]. Hashmi et al. adopted line illumination and camera-based detection reminiscent of line-scanning confocal microscopy into flow measurement of FCS in the microchannel. The multiphase flows across the entire width of the microchannel were visualized with a high-speed camera and quantified with FCS. It was demonstrated that the flow velocity of particles in the microchannel at a speed up to the order of 1 cm/s can be resolved with the technique [58]. Another approach to measure the flow directions proposed by Brinkmeier et al. was to employ two-beam FCCS instead of classical single-beam FCS into the channel [6, 59]. With two-beam FCCS, fluorescent molecules successively pass the two spatially separated confocal volumes and then the signals from two volumes are cross-correlated. The maximum in a cross-correlation curve corresponds the time that it takes for fluorophore to transport from the first to the second volume. Thus, the migration velocity and direction of fluorescent molecules between two focal volumes can be measured [6, 34, 59, 60]. C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Microfluidics and Miniaturization
2273
The dual-focus FCS (2fFCS) was developed by Enderlein and co-workers [31], and it was introduced to map the 3D flow in the microchannel [33]. In contrast to two-beam FCCS, 2fFCS makes use of the intentional overlap of the two detection regions. Several experiments demonstrated its significant advantages (freedom from calibration and increased accuracy) in the measurement of flow velocities over conventional single-focus FCS [33, 34]. An important application of flow velocity measurement based on these two-beam FCCS or based on two-focus FCCS is the measurement of fast reaction kinetic parameters. Dittrich et al. developed a spatial two-beam FCCS method to accurately determine the reaction time by measuring the exact flow time from the mixing point to the observation point in a miniaturized continuous-flow (CF) reactor. Using this CF device, the kinetics of the irreversible cleavage of a DNA strain oligomer by the enzyme exonuclease was studied [61]. It is very important to converse spatial information into the respective temporal information based on the measured flow velocity by FCCS method in the investigation on the reaction kinetics. The similar approach of “position-to-time conversion” based on FCS method was also reported by Wunderlich et al. [62]. Using this approach, single-molecule kinetics of B-domain of protein A in the microfluidic mixer devices was recorded on timescales from milliseconds to minutes.
4.2.3 Flow profiling in the interface In the proximity of an interface, the normal resolution can be significantly increased using TIR microscopy. In order to overcome the disadvantage of one-beam FCS, two-beam FCCS, or line-scan FCS on flow profiling in the close proximity of an interface caused by the limited resolution of these techniques, new TIR-FCS or TIR-FCCS configurations were developed to measure velocity profiles in the interface [36–38]. For example, Yordanov et al. tuned the evanescent wave penetration depth of TIR and the velocity at different distances less than 200 nm from the interface was measured [36]. It should be pointed out that the evanescent wave illumination is one of plane illumination methods. So by adopting the TIR illumination into FCS, more comprehensive information about the flow profile can be obtained. Another illumination configuration similar to TIR-FCS can be achieved by adopting single-plane illumination (SPIM) into FCS for measurement on the flow. Wohland et al. applied SPIM as excitation source and an electron multiplying CCD (EMCCD) as array detectors, as shown in Fig. 9 [63]. Using the technique, blood flow profiles and flow directions within living zebrafish can be determined. The most importance is that the measurement depth can be adjusted in SPIM-FCCS.
4.3 Multicomponent analyses in the channel FCS is a highly sensitive and rapid single-molecule detection technique that is often used to perform multicomponent www.electrophoresis-journal.com
2274
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
Figure 9. SPIM-FCS measurements within a live zebrafish 48 h postfertilization. (A) The ACFs of 0.2 m microspheres within a blood vessel. (B) This figure shows cross-correlation functions between the central pixel of the 20 × 20 pixel region of interest and the surrounding pixels at a distance of three pixels (same experiments as A). Reprinted with permission from [63].
analysis in static solutions based on the different diffusion times of the species through the tightly focused detection volume, which scales with the cubic root of the molecular size. In order to differentiate species based on their diffusion time, a relative difference in diffusion time between two different species of at least a factor of two is required [22]. However, in many multicomponent analyses, molecular size of the analyte species is comparable. So, it is necessary to develop some methods to enhance their difference of characteristic diffusion time in focal volume. A electrophoretic mobility shift assay method by combining FCS with CE (FCS-CE) developed by Van Orden and Keller can resolve the problem above [24]. In the method, transport of mixture of R6G and R6G-labeled dCTP (R6GdCTP) nucleotides in the capillary is measured as model by FCS when they continuously flow through the capillary in the presence of an electric field. The difference of transit times between observed R6G and R6G-dCTP in the volume increase due to their different electrical mobility in the capillary. The increased difference of transit times can agree with the accuracy requirement for two component fitting procedure in FCS measurement. The multicomponent analysis procedure based on FCS-CE is accomplished to measure the relative concentration of R6G and R6G-dCTP mixture although their molecular weight difference is minor. This method combines the selectivity of CE with the sensitivity and rapid analysis times of FCS. Compared with conventional CE method, less samples volume and short analysis time are required. The assay method was extended to investigate the interaction between DNA-protein [64, 65]. LeCaptain et al. studied the interaction of single-stranded DNA-binding protein (SSB) with fluorescently labeled 39-mer single-stranded DNA (ssDNA) [64]. It was verified that the determined binding ratios agreed with the results obtained by CE separation. The resolution of the bound and unbound complexes was significantly enhanced compared to conventional FCS. Yeh et al. combined the technique with microfluidic chip electrophoresis and study the dissociation of transcription factor Sp1DNA complex by doxorubicin [65]. The binding fractions of Sp1-DNA complex at different Sp1 concentration was measured with two-component fitting procedure of FCS. The binding constant measured by FCS was found to be comparable to that measured by conventional electrophoresis assay.
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
In comparison to FCS-CE, the coupling of twobeam FCCS with microchannel electrophoresis (two-beam FCCS/CE) shows more advantage on the multicomponent analysis. For example, in the two-beam FCCS/CE, the obtained cross-correlation function contains separate peaks corresponding to the electrophoretic flow velocities of ssDNA and conjugates of SSB with ssDNA. So the relative concentrations of the bound and unbound analytes could be determined from the curve with two component analysis [25]. Multicomponent analysis procedure based on two-beam FCCS/CE has been applied for simultaneous measurement of positive and negative ions [66] and assay of protein digests [67]. The transport of positive (cationic R6G) and negative ions (anionic 5-carboxytetramethylrhodamine) in the electrophoresis capillary was monitored by two-beam FCCS/CE [66]. The obtained cross-correlation multicomponent analysis results demonstrate that the determined relative concentrations of each analyte agreed with the expected values. Brister and Weston employed two-beam FCCS to characterize the migration rates of fluorescently labeled protein digests in a microfluidic chip electrophoresis system [67]. Meanwhile, Monte Carlo simulation methods were presented to understand the capabilities and limitations of the two-beam FCCS/CE method. The experimental and simulation results demonstrated that the application of two-beam FCCS in the electrophoresis assay can aid proteomics research significantly. In these two beam-FCCS measurements, crosscorrelation curves are obtained when fluorescent molecules successively pass the two focal volumes. However, many of the detected fluorescent molecules by the first focal volume diffuse away at the second focal volume sites and hereby cannot be measured because the focal volumes are far less than the width of channel. So only part of molecules that transit both detection regions contribute to the cross-correlation peak of two beam-FCCS, which is adverse for the accumulation of signal and detection for target molecules. In order to overcome the disadvantage, a two-beam line FCCS configure was developed by Schiro et al. for use in continuous-flow single-molecule CE, where two rectangular illumination volumes completely match with the width of the microchannel, as shown in Fig. 10 [68]. The combination of this configure with CE was used to determine the mixture of FITC, FITC-labeled glycine, and FITC-labeled glutamate. In the
www.electrophoresis-journal.com
Microfluidics and Miniaturization
Electrophoresis 2014, 35, 2267–2278
Figure 10. (A) Schematic illustrating the experimental setup for continuous-flow single-molecule CE. (B) Simulation showing the distribution of laser intensity across the width of a 2-m channel for two spatially separated focal volumes. Reprinted with permission from [68].
geometry, it was observed that nearly every molecule in a sample was detected. This high detection efficiency is critical for the chemical analysis on low copies of molecules in single cells or subcellular compartments.
4.4 DNA analysis Combination of FCS with microchannel electrophoresis shows great potential in the study of fundamental physical properties of DNA undergoing electrophoretic migration [69, 70]. For example, it was observed that the electrophoretic behavior of DNA in the microchannel was highly dependent upon the electric field as the migration speed linearly increases with the applied electric field [69]. When FCCS was used to study electrophoretic behavior of DNA in the microchannel, both the translational diffusion and electrophoretic flow properties could be obtained from the measurement. For example, electrophoretic mobilities of different length of double-stranded DNA (75–1019 bp) in microfluidic channels were determined using dfFCS. It was found that when dsDNA were separated in the free solution, their mobility is independent of DNA length. However, when polyethylene oxide was sieved as the separation media, the electrophoretic rates become size dependent with a power-law exponent between 0.28 and 0.31 [70]. The role of counterions in modulating the effective charge and diffusion properties of ssDNA was investigated by C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
2275
Fogarty et al. [71]. It was observed that with the concentration increase of Mg2+ , the migration velocity of ssDNA decreased but the rate of ssDNA translational diffusion increased. The results that larger diffusion constants and smaller hydrodynamic radii indicated that the existence of Mg2+ neutralized the negative charges on the ssDNA backbone, which lowers the self-repulsion between nucleotides and makes the ssDNA form more compact structures. The kinetics and energetics of DNA hairpin folding was investigated by Jung et al. using simultaneous two-beam FCCS and single-beam FCS [26,72]. In the hairpin DNA, each end is attached with a dye and a quencher. The folding and unfolding of hairpin DNA results in intensity fluctuation due to the fluorescence on/off characteristic caused by the contact between the fluorophor and the quencher. The correlation function determined with FCS thus contains fluctuation contributions characterized by a diffusional relaxation time, d , and a relaxation time for conformational fluctuations, R . But it is difficult for conventional FCS to resolve two relaxation times because the relaxation process occurs on a time scale similar to or less than the sampling interval. Jung et al. successfully decoupled the different relaxation times of hairpin DNA by analyzing fluorescence fluctuations of hairpin DNA in a capillary observed at two spatially offset detection volumes with two-beam FCCS. The results showed a three-state mechanism was involved in the DNA hairpin folding reaction and a long-lived intermediate form of the DNA hairpin was included in the reaction.
4.5 Characterization on some parameters of nanoparticles Ren and co-workers developed a method by combining FCS with microfluidic chip electrophoresis to measure the surface charge of aqueous mercaptopropionic acid (MPA)-coated cadmium telluride (CdTe) QDs as shown in Fig. 3 [28]. According to the Stokes–Einstein equation, diffusion coefficients of spherical particles, D are inversely proportional to its hydrodynamic radius, R as indicated in Eq. (9): D=
kT , 6R
(9)
where kT is the thermal energy and is the viscosity of the solution. Thus, according to Eqs. (3) and (9), the hydrodynamic radius, R of QDs can be calculated as indicated in Eq. (10): R=
2kT D . 302
(10)
As shown in Eq. (7), migration velocity, Vf of QDs in the channel can be calculated based on the parameters (0 and f ) by fitting FCS raw data with Eq. (6). QDs migrates in the microfluidic channel with no electrosmotic flow under the electric field, so their migration velocity Vf is positively www.electrophoresis-journal.com
2276
C. Dong and J. Ren
Electrophoresis 2014, 35, 2267–2278
tures. It is expected that the approach can be straightforwardly extended to simultaneous detection of several virus types.
5 Conclusion
Figure 11. Normalized FCS curves of aqueous CdTe QDs in the microchannel under the different electric field strength. Reprinted with permission from [28].
proportional to the intensity of electric field E and can be expressed as indicated in Eq. (11): q E, (11) Vf = 6R where q is the surface charge of fluorescent particles, and E the intensity of electric field. So, when R in Eq. (11) is replaced with one in Eq. (10) and Eq. (11) is combined with Eq. (7), the relationship between the average migration time f and the intensity of electric field E can be deduced as Eq. (12): 0 q 1 = E = s × E. f 4kT D
(12)
As a result, the charge q can be calculated on the slope (s) from the linear curve of 1/ f versus E (q = 4ksT D /0 ). Figure 11 is the normalized fluorescence autocorrelation function of QDs under the different electric field strength. It demonstrated that the transit time decreased with the increased electric field strength. The measured results show that the charges of QDs increased with pH value of solution. When surface ligand-mercaptopropionic acid was replaced by glutathione, the surface charge of CdTe QDs was increased by about 55.3%. This result showed that the surface charge of QDs was remarkably associated with the type of stabilizers on QDs surface, buffer pH, and other factors.
The applications of combination of single-molecule detection technique—FCS with channel showed more and more advantages over the sole channel separation methods. (i) Lower sample consumption. The technique of combination of FCS with channel assay has extremely high sensitivity. Single molecules are detected when they are separated in the channel. Herein, it can lower the requirement for sample volume and concentration. It is critical for the biochemical assay of low copies of molecules in single cells. (ii) More temporal information. Compared with the conventional channel separation method such as microfluidic chip electrophoresis, implementation of FCS in channel can provide the dynamics information in the time scale from microsecond to second such as the microsecond relaxation and translation of molecules. (iii) Enhanced spatial information. Narrow channels are more regarded as a miniaturized reaction and separation platform. The combination of FCS can provide the 3D spatial information about reaction and separation in the channels. It can be concluded that the combination strategies is emerging but important in the analytical chemistry, analytical biochemistry, high-throughput drug screening, and in vivo study of living cells. This work was financially supported by NFSC (20905048, 21075081, 21135004, and 21327004), Innovation Program of Shanghai Municipal Education Commission (14ZZ024), Shanghai Educational Development Foundation (2008CG12), and SMC-Chenxin Young Scholar project sponsored by Shanghai Jiao Tong University. The authors have declared no conflict of interest.
6 References [1] Punj, D., Mivelle, M., Moparthi, S. B., Van Zanten, T. S., Rigneault, H., Van Hulst, N. F., Garcia-Parajo, M. F., Wenger, J., Nat. Nanotechnol. 2013, 8, 512–516. [2] Kim, S. A., Heinze, K. G., Schwille, P., Nat. Methods 2007, 4, 963–973. [3] Zamir, E., Lommerse, P. H. M., Kinkhabwala, A., Grecco, H. E., Bastiaens, P. I. H., Nat. Methods 2010, 7, 295–298.
4.6 Detection of viruses Rudenko et al. demonstrate the possibility to detect the Q bacteriophage on the single-virus level by integrating microfluidic chip with FCS system [73]. Multiparameters on viruses including diffusion coefficient, flow velocity, and concentration of viruses were extracted from FCS experiments. Using the approach, the signal of single phage can be distinguished from their structural parts, agglomerates, or mix C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
[4] Magde, D., Elson, E. L., Webb, W. W., Phys. Rev. Lett. 1972, 29, 705–708. ¨ Widengren, J., Kask, P., Eur. Biophys. [5] Rigler, R., Mets, U., J. 1993, 22, 169–175. [6] Brinkmeier, M., Dorre, K., Stephan, J., Eigen, M., Anal. Chem. 1999, 71, 609–616. [7] Rigler, R., Biochem. Biophys. Res. Commun. 2010, 396, 170–175. [8] Elson, E. L., Biophys. J. 2011, 101, 2855–2870.
www.electrophoresis-journal.com
Electrophoresis 2014, 35, 2267–2278
Microfluidics and Miniaturization
2277
[9] Ruan, L. G., Xu, Z. C., Lan, T., Wang, J. J., Liu, H., Li, C. D., Dong, C. Q., Ren, J. C., Anal. Chem. 2012, 84, 7350–7358.
[36] Yordanov, S., Best, A., Butt, H. J., Koynov, K., Opt. Express 2009, 17, 21149–21158.
[10] Dong, C. Q., Chowdhury, B., Irudayaraj, J., Analyst 2013, 138, 2871–2876.
[37] Blom, H., Hassler, K., Chmyrov, A., Widengren, J., Int. J. Mol. Sci. 2010, 11, 386–406.
[11] Dong, C. Q., Qian, H. F., Fang, N. H., Ren, J. C., J. Phys. Chem. B 2006, 110, 11069–11075.
[38] Schmitz, R., Yordanov, S., Butt, H. J., Koynov, K., Dunweg, B., Phys. Rev. E 2011, 84, 066306.
[12] Dong, C. Q., Qian, H. F., Fang, N. H., Ren, J. C., J. Phys. Chem. C 2007, 111, 7918–7923.
[39] Liao, D., Galajda, P., Riehn, R., Ilic, R., Puchalla, J. L., Yu, H. G., Craighead, H. G., Austin, R. H., Opt. Express 2008, 16, 10077–10090.
[13] Tang, L. C., Dong, C. Q., Ren, J. C., Talanta 2010, 81, 1560–1567. [14] Manz, A., Graber, N., Widmer, H. M., Sens. Actuators B. Chem. 1990, 1, 244–248. [15] Harrison, D. J., Manz, A., Fan, Z., Luedi, H., Widmer, H. M., Anal. Chem. 1992, 64, 1926–1932.
[40] Selmke, M., Schachoff, R., Braun, M., Cichos, F., RSC Adv. 2013, 3, 394–400. [41] Gennerich, A., Schild, D., Biophys. J. 2000, 79, 3294–3306.
[16] Harrison, D. J., Fluri, K., Seiler, K., Fan, Z., Effenhauser, C. S., Manz, A., Science 1993, 261, 895–897.
[42] 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.
[17] Arora, A., Simone, G., Salieb-Beugelaar, G. B., Kim, J. T., Manz, A., Anal. Chem. 2010, 82, 4830–4847.
[43] Petrasek, Z., Krishnan, M., Monch, I., Schwille, P., Microsc. Res. Techniq. 2007, 70, 459–466.
[18] Kovarik, M. L., Ornoff, D. M., Melvin, A. T., Dobes, N. C., Wang, Y., Dickinson, A. J., Gach, P. C., Shah, P. K., Allbritton, N. L., Anal. Chem. 2013, 85, 451–472.
[44] Sanguigno, L., De Santo, I., Causa, F., Netti, P., Anal. Chem. 2010, 82, 9663–9670.
[19] Dittrich, P. S., Manz, A., Nat. Rev. Drug Discov. 2006, 5, 210–218.
[46] Meinhart, C. D., Wereley, S. T., Santiago, J. G., Exp. Fluids 1999, 27, 414–419.
[20] Fogarty, K., Van Orden, A., Methods 2009, 47, 151–158. [21] Dong, C. Q., Ren, J. C., Analyst 2010, 135, 1395–1399.
[47] Devasenathipathy, S., Santiago, J. G., Takehara, K., Anal. Chem. 2002, 74, 3704–3713.
[22] Schwille, P., Meyer-Almes, F. J., Rigler, R., Biophys. J. 1997, 72, 1878–1886.
[48] Gai, H. W., Li, Y., Silber-Li, Z., Ma, Y. F., Lin, B. C., Lab Chip 2005, 5, 443–449.
[23] Magde, D., Web, W. W., Elson, E. L., Biopolymers 1978, 17, 361–367.
[49] Edel, J. B., Hill, E. K., de Mello, A. J., Analyst 2001, 126, 1953–1957.
[24] Van Orden, A., Keller, R. A., Anal. Chem. 1998, 70, 4463–4471.
[50] Liu, T. C., Wohland, T., Watt, F., Bettiol, A., Udalagama, C., Wang, L. P., Med. Phys. 2012, 39, 5060–5064.
[25] LeCaptain, D. J., Van Orden, A., Anal. Chem. 2002, 74, 1171–1176.
[51] Brister, P. C., Kuricheti, K. K., Buschmann, V., Weston, K. D., Lab Chip 2005, 5, 785–791.
[26] Jung, J., Van Orden, A., J. Am. Chem. Soc. 2006, 128, 1240–1249.
[52] Kuricheti, K. K., Buschmann, V., Weston, K. D., Appl. Spectrosc. 2004, 58, 1180–1186.
¨ [27] Gosch, M., Blom, H., Holm, J., Heino, T., Rigler, R., Anal. Chem. 2000, 72, 3260–3265.
[53] Kunst, B. H., Schots, A., Visser, A. J. W. G., Anal. Chem. 2002, 74, 5350–5357.
[28] Dong, C. Q., Bi, R., Qian, H. F., Li, L., Ren, J. C., Small 2006, 2, 534–538.
[54] Lenne, P. F., Colombo, D., Giovannini, H., Rigneault, H., Single Mol. 2002, 3, 194–200.
[29] Chen, L., Ren, J. C., Bi, R., Chen, D., Electrophoresis 2004, 25, 914–921.
[55] Park, H. Y., Qiu, X. Y., Rhoades, E., Korlach, J., Kwok, L. W., Zipfel, W. R., Webb, W. W., Pollack, L., Anal. Chem. 2006, 78, 4465–4473.
[30] Foquet, M., Korlach, J., Zipfel, W. R., Webb, W. W., Craighead, H. G., Anal. Chem. 2004, 76, 1618–1626. [31] Dertinger, T., Pacheco, V., von der Hocht, I., Hartmann, R., Gregor, I., Enderlein, J., Chemphyschem 2007, 8, 433–443. [32] Felten, M., Staroske, W., Jaeger, M. S., Schwille, P., Duschl, C., Electrophoresis 2008, 29, 2987–2996. [33] Arbour, T. J., Enderlein, J., Lab Chip 2010, 10, 1286–1292.
[45] De Santo, I., Causa, F., Netti, P. A., Anal. Chem. 2010, 82, 997–1005.
[56] Liu, K., Tian, Y., Burrows, S. M., Reif, R. D., Pappas, D., Anal. Chim. Acta 2009, 651, 85–90. [57] Carlotto, S., Fortunati, I., Ferrante, C., Schwille, P., Polimeno, A., Microfluid Nanofluid 2011, 10, 551–561. [58] Hashmi, S. M., Loewenberg, M., Dufresne, E. R., Opt. Express 2007, 15, 6528–6533. [59] Brinkmeier, M., Dorre, K., Riebeseel, K., Rigler, R., Biophys. Chem. 1997, 66, 229–239.
[34] Lumma, D., Best, A., Gansen, A., Feuillebois, F., Radler, J. O., Vinogradova, O. I., Phys. Rev. E 2003, 67, 056313.
[60] Dittrich, P. S., Schwille, P., Anal. Chem. 2002, 74, 4472–4479.
[35] Pan, X. T., Yu, H., Shi, X. K., Korzh, V., Wohland, T., J. Biomed. Opt. 2007, 12, 014034.
[61] Dittrich, P. S., Muller, B., Schwille, P., Phys. Chem. Chem. Phys. 2004, 6, 4416–4420.
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
www.electrophoresis-journal.com
2278
C. Dong and J. Ren
[62] Wunderlich, B., Nettels, D., Benke, S., Clark, J., Weidner, S., Hofmann, H., Pfeil, S. H., Schuler, B., Nat. Protoc. 2013, 8, 1459–1474. [63] Wohland, T., Shi, X. K., Sankaran, J., Stelzer, E. H. K., Opt. Express 2010, 18, 10627–10641. [64] LeCaptain, D. J., Michel, M. A., Van Orden, A., Analyst 2001, 126, 1279–1284. [65] Yeh, H. C., Puleo, C. M., Lim, T. C., Ho, Y. P., Giza, P. E., Huang, R. C., Wang, T. H., Nucleic Acids Res. 2006, 34, e144. [66] Fogarty, K., Van Orden, A., Anal. Chem. 2003, 75, 6634–6641. [67] Brister, P. C., Weston, K. D., Analyst 2006, 131, 303–310.
C 2014 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Electrophoresis 2014, 35, 2267–2278
[68] Schiro, P. G., Kuyper, C. L., Chiu, D. T., Electrophoresis 2007, 28, 2430–2438. [69] Foquet, M., Korlach, J., Zipfel, W., Webb, W. W., Craighead, H. G., Anal. Chem. 2002, 74, 1415–1422. [70] Bayer, J., Radler, J. O., Electrophoresis 2006, 27, 3952–3963. [71] Fogarty, K., McPhee, J. T., Scott, E., Van Orden, A., Anal. Chem. 2009, 81, 465–472. [72] Jung, J., Ihly, R., Scott, E., Yu, M., Van Orden, A., J. Phys. Chem. B 2008, 112, 127–133. [73] Rudenko, M. I., Kuhn, S., Lunt, E. J., Deamer, D. W., Hawkins, A. R., Schmidt, H., Biosens. Bioelectron. 2009, 24, 3258–3263.
www.electrophoresis-journal.com
