Article pubs.acs.org/ac

Planar Electrochromatography Using an Electrospun Polymer Nanofiber Layer Toni E. Newsome and Susan V. Olesik* Department of Chemistry and Biochemistry, The Ohio State University, 100 West 18th Avenue, Columbus, Ohio 43210, United States S Supporting Information *

ABSTRACT: Electrospun polymer nanofiber stationary phases were examined for their application to planar electrochromatography (PEC). Separations were performed on polyacrylonitrile nanofiber ultra-thin-layer chromatography (UTLC) plates in 1−2 min using a ternary mobile phase. The influences of buffer concentration and pH, ratio of organic modifier, and development time on analyte migration distances were studied. Band broadening in this system was studied as a function of distance. The plate height initially decreased and then plateaued with a minimum plate height value as low as 11 μm. Nanofiber alignment considerably increased analyte migration rate, but larger spot sizes were noted when nearly complete fiber alignment was used. The easily tunable stationary phase thickness can be tailored to a given separation, where thinner layers promote faster separations and thicker layers are ideal for more complex mixtures. Compared to UTLC, PEC offers unique selectivity and decreased analysis time (>4 times faster over 15 mm). Results for a two-dimensional separation using UTLC and PEC are also reported. These rapid separations required 11 min using a 40 × 40 mm plate and exhibited a significant increase in separation number (70−77).

T

heat produced during PEC leads to two disadvantages: evaporation of mobile phase and flux of mobile phase to the stationary phase surface.5,8 The former leads to poor repeatability, while the latter leads to considerable band broadening.8 These effects can be partially eliminated by covering the layer but can be fully eliminated by pressurizing the layer, i.e., pressurized PEC (PPEC).5,9,12−14 Refinement of PPEC continues;4,7,9,10,15−18 however, these devices have yet to be commercialized, perhaps due to their complexity.19 Another challenge is the production of stationary phases designed for PPEC.4,5,8 So far, commercial TLC and HPTLC plates have been applied. However, the relatively large thicknesses of these plates make them ineffective for dissipating the Joule heat produced. To date, Nurok and coworkers exclusively have studied thinner layers dedicated to PPEC.7 While only preliminary results have been reported, the authors stated that the 125 μm thick monolithic polymer layer holds promise for fast separations of biological compounds.4,7 Further development of PPEC certainly requires plates with thinner layers than commercial plates, which are more appropriate for heat dissipation.4,8 Electrospun nanofiber stationary phases were recently introduced for UTLC, in a technique referred to as E-UTLC.20 Electrospinning is a process that relies on repulsive electrostatic forces to produce nanofibers from a polymer solution.21 In EUTLC, electrospinning is used to fabricate a mat of randomly placed nanofibers ∼25 μm thick which is used as a stationary phase for UTLC. E-UTLC plates offer many advantages, including

hin-layer chromatography (TLC), or planar chromatography, remains a widely used technique in synthetic chemistry and in various industries.1 High-performance thin-layer chromatography (HPTLC) and ultra-thin-layer chromatography (UTLC) have also been developed.2 HPTLC offers shorter analysis times and lower detection limits by utilizing sorbent particles with smaller dimensions and size distributions. UTLC further decreases analysis time by employing layers with thicknesses of 5−25 μm (as opposed to TLC with thicknesses of 100−400 μm).2,3 The main disadvantage of conventional TLC is the capillary-driven, nonconstant mobile phase flow.4,5 The velocity decreases with increasing distance, so it cannot be optimized to minimize band broadening (or plate height, H) as is done in gas chromatography and high-performance liquid chromatography.2,4 Consequently, the minimum plate height (Hmin) for TLC depends on the position of the analyte on the plate, and H significantly increases at distances past Hmin as mobile phase velocity diminishes considerably.2,6 While HPTLC and UTLC have shorter separation times, use of these phases with conventional capillary-driven mobile phases limits the peak capacity which can be achieved.7 Consequently, forced-flow techniques have been developed to eliminate drawbacks of capillary flow, including rotational planar chromatography, overpressure layer chromatography, and planar electrochromatography (PEC).2,7 In PEC, the mobile phase is driven by the electroosmotic effect by applying a potential across the stationary phase.4,8−11 Compared to TLC, the electroosmotically driven mobile phase velocity is constant and independent of separation distance and particle size (within a restricted range) under constant conditions.5,8 Important advantages over conventional TLC include higher speed of separation and performance.8,11 However, Joule © 2014 American Chemical Society

Received: September 23, 2014 Accepted: October 4, 2014 Published: October 21, 2014 10961

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

easily tunable thicknesses and nanofiber diameters.20,22 In addition, rapid separations and enhanced efficiencies have been reported for trials using short separation distances (15−30 mm). This method is capable of producing nanofibers with diverse functionality.21 So far, polymer,20,23−25 carbon,22 and silica26 nanofibers have been utilized. Polymer stationary phases have been produced from polyacrylonitrile (PAN),20,23−25 poly(vinyl alcohol),27 and cellulose acetate.28 Randomly placed and highly aligned nanofibers were compared as stationary phases in E-UTLC; the aligned fibers provided increased speed of analysis and efficiency compared to randomly placed nanofibers.23,26 Although faster separations and enhanced efficiencies are possible with E-UTLC, the mobile phase velocity nevertheless diminishes as a function of distance since the flow is capillarydriven when using traditional methods.20,22−27 Therefore, the performance of electrospun plates may be further improved by using forced-flow-driven mobile phases. Furthermore, poor selectivity has been noted for a set of laser dyes using PAN nanofiber plates because analytes in this mixture have very similar chemical structures.20,23,25 Because of the contribution of electrophoresis to the separation mechanism, it is expected that PEC can offer additional selectivity relative to UTLC, as the analytes are composed of charged and neutral species.5 Additionally, the thin and tunable thickness of electrospun stationary phases offers a promising solution to problems associated with Joule heating in thicker layers currently used in PPEC. Accordingly, herein the performance of electrospun nanofiber stationary phases is examined using PEC and twodimensional (2D) UTLC-PEC separation techniques.

when the lid was removed, so that it acted as a safety interlock to protect the operator from exposure to electrical current. The HVPS was turned on when the mobile phase fronts met each other at the origin. A current of 1 kV was applied across the plate, and separations were performed for 1 min using randomly placed PAN nanofibrous stationary phases electrospun with a 20 min collection time, unless otherwise noted. After electrospun PAN nanofiber stationary phases were prepared, 50 nL of each analyte solution was spotted onto the plates using capillary tubing (100 μm i.d., 200 μm o.d., Polymicro Technologies, Phoenix, AZ). The analyte origin was 1.0 cm from the plate edge closest to the anode. The laser dye concentrations in methanol were 5 × 10−5 M for kiton red and the three rhodamine dyes, 10−4 M for sulforhodamine, and 10−3 M for pyrromethene. A 20 mL portion of mobile phase was used in each reservoir. Wicks were made from Whatman 3MM paper. A current of 1 kV was applied in constant voltage mode, and separations were performed for 1 min unless noted otherwise. Visualization was done using UV light (λ = 254 nm) and a Canon A650IS digital camera mounted to a documentation system (Spectroline, Westbury, NY).20,23,26 Measurements were performed using ImageJ (http://rsbweb.nih.gov/ij/). Reported results were based on at least three measurements. Separations were performed on randomly placed nanofibrous stationary phases which were electrospun for 20 min unless otherwise noted. Ultra-Thin-Layer Chromatography and Two-Dimensional Separations. After nanofiber stationary phases were prepared, each analyte was spotted onto plates for UTLC similar to PEC. Unlike PEC, the position of the analyte origin was 0.5 cm from the edge of the plate in contact with the mobile phase. Specifications for UTLC can be found in the SI. Onedimensional UTLC was performed on 2.5 × 5.0 cm plates, and 2D separations were performed on 4.0 × 4.0 cm plates. For 2D UTLC-PEC, UTLC was performed in the first dimension as described above, the stationary phase was dried and transferred to a glass substrate, and PEC was performed in the second dimension as described in Planar Electrochromatography.

■

MATERIALS AND METHODS Materials. Polyacrylonitrile (PAN), average Mw ≈ 150 000, was purchased from Sigma-Aldrich (St. Louis, MO). N,NDimethylformamide (99.9%), citric acid, acetonitrile (ACN), 2-propanol (2-PrOH), and methanol (MeOH, ACS grade) were purchased from Fisher Scientific (Pittsburgh, PA). Laser dyes were purchased from Exciton Inc. (Dayton, OH); these included kiton red 620 (KR), pyrromethene 597 (PM), rhodamine 101 (R101), rhodamine 590 chloride (R590), rhodamine 610 chloride (R610), and sulforhodamine 640 (SR). Sodium citrate was purchased from Jenneile Chemical (Cincinnati, OH). Planar Electrochromatography. Preparation of electrospun stationary phases is described in the Supporting Information (SI). A PEC apparatus was assembled from an electrophoresis chamber (Mini-Sub Cell GT, Bio-Rad, Hercules, CA) and a highvoltage power supply (HVPS; FisherBiotech Electrophoresis Power Supply, FB1000, 1 kV, Pittsburgh, PA). The details of the apparatus adapted for PEC are shown in Figure S-1 in the SI. Compared to UTLC, slight modifications in the preparation of the stationary phases were adapted for PEC, namely the position of the analyte origin and pre-wetting of the stationary phase with mobile phase. Pre-wetting was achieved by soaking Whatman 3MM paper (Sigma-Aldrich) in mobile phase for several minutes, and the stationary phase was dampened with the paper on both sides of the analyte origin. Electrical connections were made between the mobile phase and stationary phase using the Whatman paper as a wick. Each wick overlapped the stationary phase by 0.5 cm on each side of the plate (SI, Figure S-2). This provided 0.5 cm between the origin and wick on the anode side and 3.5 or 2.5 cm (depending on plate size, Figure S-2, parts A and B, respectively) between the origin and wick on the cathode side. A glass cover plate was placed on the stationary phase, and the chamber lid was attached. Current to the chamber was broken

■

RESULTS AND DISCUSSION Initial Binary Mobile Phase Separations. Randomly placed and aligned PAN nanofiber stationary phases were electrospun for PEC in a similar manner to those for UTLC.20,23 The separation of these laser dyes using electrospun PAN stationary phases was not achieved previously using UTLC.20,23 This separation is difficult because the dyes have very similar chemical structures (SI, Table S-1). For example, R590 and R610 differ only by two ethyl groups and the carboxyl functional group. By incorporating electrophoretic effects into the separation, PEC should offer distinctly different selectivity to aid in the separation of these analytes relative to UTLC since the dyes vary in charge. First, the PEC separation of four laser dyes, KR, R590, R610, and SR, was attempted using binary mobile phases comprised of organic solvent with aqueous citrate buffer. Previous studies on the separation of these laser dyes showed that mobile phases with ACN or 2-PrOH mixed with aqueous buffer provide adequate selectivity.14 PEC was performed for 1 min using 50:50 ACN/10 mM citrate buffer, pH 6.0 (v/v), and 50:50 2-PrOH/25 mM citrate buffer, pH 6.0 (v/v). Analyte migration distances are compared in Table 1. Under these conditions, electroosmotic flow velocity, υeo, was faster than electrophoretic velocity of the ions, υep, since all species migrated in the same direction (toward the cathode) 10962

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

Table 1. Migration Distance of Laser Dyes Using Different Mobile Phasesa migration distance (mm) laser dye KR R590 R610 SR

ACN/10 mM buffer (50:50) 9.04 0.19 6.48 7.05

± ± ± ±

1.16 0.08 0.43 0.81

2-PrOH/25 mM buffer (50:50) 1.13 3.43 2.35 0.26

± ± ± ±

0.89 1.05 0.98 0.78

a

Conditions: potential applied for 1 min, 50:50 ACN/10 mM citrate buffer and 50:50 2-PrOH/25 mM citrate buffer, pH 6.0 (v/v), mobile phases run at 1 kV.

regardless of charge but at different rates. Retention order differed in the two mobile phases in that KR and SR, the two negatively charged analytes, were the least retained species in the ACN/buffer mobile phase and the most retained in the 2-PrOH/buffer mobile phase. The opposite was observed for positively charged R590. In electrophoretic separations, both υeo and υep are toward the cathode for cations, and for anions, υeo is toward the cathode while υep is toward the anode, resulting in the retention order of cations < neutrals < anions.29 Because the order of retention changed with different organic solvents in the mobile phase and because the negatively charged dyes were the least retained in the ACN/buffer mobile phase, electrochromatography rather than electrophoresis controls the retention order of analytes.29 Differences in selectivity were attributed to different chromatographic interactions of analytes with the organic component (ACN or 2-PrOH) of the mobile phase. In contrast, differences in analyte migrations rates were attributed to electrophoretic effects. For example, the least retained analytes tended to migrate faster in the ACN/ buffer mobile phase. The apparent higher electrophoretic mobilities, μep, observed were a result of the lower viscosity of ACN relative to 2-PrOH. Accordingly, ACN/aqueous buffer mobile phases are the most popular choice in electrochromatography.29 Even though faster migration rates using the ACN/buffer mobile phase could have led to faster separations, developed spot shapes were smaller and more reproducible in the 2-PrOH/buffer mobile phase. Poor spot shapes with the ACN/buffer mobile phase were likely caused by resistance to mass transfer at the faster migration rates. Therefore, ternary mixtures of ACN, 2-PrOH, and buffer were examined to integrate the better spot shapes using 2-PrOH with the better speed of separation using ACN. Ternary Mobile Phase Separations. Effect of Organic Solvent Concentrations. Ternary mixtures of ACN and 2-PrOH with aqueous citrate buffer were examined in the separation of six laser dyes: KR, PM, R101, R590, R610, and SR. Separations were performed for 1 min using a 50:50 ratio of total organic solvent to aqueous buffer while varying the ratio of ACN to 2-PrOH. R101 and R610 are known to exist in two principal forms, cationic or zwitterionic, attributed to protonation or deprotonation of their carboxyl groups, respectively. Under the conditions used, R101 and R610 are mostly in zwitterionic form because the carboxyl groups are deprotonated (pKa = 3.3 and 3.6, respectively).30 Both analytes increased in distance with increasing ACN concentration (decreasing 2-PrOH concentration) as the viscosity of the mobile phase decreased; their retention behavior was similar at all ratios of ACN:2-PrOH because of their similar chemical structure and charge (Figure 1A). KR and SR showed similar increases in distance with ACN concentration, and similar retention variation at all ratios of ACN:2-PrOH because of their negative charge and

Figure 1. Migration distance of laser dyes versus ACN and 2-PrOH concentrations: (A) KR (dark blue diamonds), R101 (orange circles), R610 (purple squares), and SR (light blue circles); (B) PM (red squares) and R590 (green triangles). Conditions: potential applied for 1 min, total concentration of organic (ACN plus 2-PrOH) is 50%, 50% 25 mM citrate buffer, pH 6.0, run at 1 kV.

similar chemical structure. Similarly, the neutral PM and positively charged R590 initially increased in migration with increasing ACN concentration; however, they began to decrease in migration between 30% and 35% ACN (Figure 1B). This reversed the order of retention for this group of analytes relative to the rest of the mixture, as they were the least retained at high 2-PrOH concentrations and the most retained at high ACN concentrations. In addition, R590 undergoes base hydrolysis of the ester group around pH 7.5; since the pH of the ternary mobile phase shifts upward with increasing ACN content, the charge of R590 begins to change from positive to more neutral with increasing concentration of ACN (SI, Table S-1).31−33 Therefore, the change in retention order was a combined result of the chromatographic contribution to this separation as well as the change between charged and neutral states in different mobile phase mixtures.34 Selectivity and peak shape were better at higher concentrations of 2-PrOH, while migration distances of the least retained analytes were greater at higher concentrations of ACN. To balance selectivity and speed of separation, mobile phases comprised of 25:25:50 ACN/2-PrOH/buffer (v/v/v) were selected for further experiments. Effect of Buffer Concentration. The influence of buffer concentration on analyte migration distance was explored for a 1 min separation with 25:25:50 ACN/2-PrOH/citrate buffer, pH 6.0 (v/v/v), using buffer concentrations ranging from 15 to 30 mM (SI, Figure S-3). Retention order remained the same, 10963

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

and no change in selectivity was observed. A notable increase in migration distance was observed as concentration increased (from 15 to 25 mM), and a plateau occurred at 30 mM. In addition, enhanced dispersion in the developed spots occurred for mobile phases with 15 mM buffer compared to those with higher concentrations. Theory predicts that high concentrations of buffer decrease the electrical double-layer thickness (δ),8

δ=

ε0εrRT 2cF 2

(1)

where c is the buffer concentration, ε0 is the permittivity of vacuum, εr is the dielectric constant, F is the Faraday constant, R is the gas constant, and T is temperature. According to eq 1, the rate of electroosmotic flow (EOF) should decrease with increasing ionic strength. However, previous studies on capillary electrochromatography (CEC) and PEC do not confirm this effect.8,10,16,35−39 Rather, an increase in buffer concentration leads to increased mobile phase flow rates and thus increased analyte migration distances; once the concentration has increased past a certain extent, there is decreased analyte migration. Moreover, when buffer concentrations are low, large dispersion of the developed spots was noted, even though band dispersion typically decreases with migration distances.5,37 Between 15 and 25 mM buffer concentrations, increased flow rate with buffer concentration was explained by the diminishing overlap of the electrical double layer, as the size of the double layer decreases with increasing buffer concentration.8,10,35 Joule heating leads to the evaporation of mobile phase from the stationary phase and flux of mobile phase to the surface of the stationary phase,5 and the generation of Joule heat increases with increasing buffer concentration. If the buffer concentration is too low, excess solvent created on the stationary phase surface by mobile phase flux cannot be fully evaporated, and pooling leads to dispersion of developed spots. This effect was observed at 15 mM. If concentration is too high, strong evaporation leads to dry zones in the stationary phase, which impedes analyte migration; this began to take effect at 30 mM. Therefore, appropriate buffer concentration selection can minimize the effects related to Joule heating, and moderate concentrations between 20 and 25 mM work best. The effects of Joule heat can be partially eliminated by covering the layer with a counterplate, as was done here;12,13 however, they have only been fully eliminated by using PPEC.5,9 The remaining separations utilized 25 mM buffer to achieve the highest speed of separation without encountering drying effects. Effect of Buffer pH. The impact of buffer pH on analyte migration distance was explored (Figure 2) using 25 mM citrate buffers ranging in pH between 4.0 and 6.0. The change in migration for the uncharged analyte, PM, reflects the change in rate of EOF, and its migration distance increased with pH, which is typical in electrochromatographic separations with silica stationary phases. As pH increases, there is an increase in charge density of the electrical double layer at the stationary phase−mobile phase interface, and thus an enhancement in the rate of EOF. As discussed in Effect of Organic Solvent Concentrations, R101 and R610 exist mostly in zwitterionic form at pH 6.0. As pH decreased and approached the pKa of the dyes, an increasing portion of each analyte also existed in cationic form. Therefore, as pH decreased, R01 and R610 began to migrate farther (in the direction of the cathode) relative to neutral PM and eventually had similar retention

Figure 2. Migration distance of laser dyes versus buffer pH. Analytes: KR (dark blue diamonds), PM (red squares), R101 (orange circles), R590 (green triangles), R610 (purple squares), and SR (light blue circles). Conditions: potential applied for 1 min, 25:25:50 ACN/ 2-PrOH/25 mM buffer (v/v/v), run at 1 kV.

compared to positively charged R590. The effect was more dramatic with R610, which was expected to have a greater percentage in cationic form because of its higher pKa relative to that of R101. As pH decreased, the initially negatively charged KR and SR also began to migrate farther as the amine groups become protonated.40 To achieve the highest selectively, the remaining separations utilized mobile phases comprised of 25 mM citrate buffer at pH 5.6. Effect of Development Time. The impact of development time on migration distance was investigated (Figure 3). When

Figure 3. Migration distance of laser dyes versus separation time. Analytes: KR (dark blue diamonds), PM (red squares), R101 (orange circles), R590 (green triangles), R610 (purple squares), and SR (light blue circles). Conditions: 25:25:50 ACN/2-PrOH/25 mM citrate buffer, pH 5.6 (v/v/v), run at 1 kV.

the HVPS is turned on at the beginning of the separation, about 15 s elapses before the system reaches 900 V, and an additional 10−15 s is needed to fully reach 1 kV. Therefore, times of 1 min or more were chosen so that the separation was under a constant 1 kV potential for at least 30 s. Initially, migration distance was linearly dependent on separation time between 60 and 90 s, indicating that operating conditions remained relatively constant in this system.9 However, there was a notable increase in migration rate (5−150%) when the time was increased to 120 s, suggesting that conditions changed during the separation to enhance EOF. Joule heating caused the temperature of the PEC system to increase at this separation time, as noted by the formation of condensate on the chamber 10964

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

lid. Temperature has a large influence on the retention and quality of a separation, and the increased migration distances at 2 min were attributed to the increase in EOF resulting from the temperature increase according to eq 1.8,16 Other covered PEC systems under minimal pressure have shown a similar increase in migration rate with separation time due to increasing temperature.15 In CEC, the effects of Joule heating are controlled by the heat dissipation of the capillary walls possessing high surface area-to-volume ratio. Nurok et al.37 also noted that, in PEC, operation under constant power conditions instead of constant voltage stabilized the system to provide constant power and Joule heating. With PPEC, the relatively large thickness of the TLC stationary phase is less effective at dissipating this heat.8 However, often an alumina nitride ceramic is used in PPEC to minimize the impact of Joule heating to maintain the linearity of migration distance with time.9 Pressurizing the system helps control these effects, and the use of thinner planar stationary phases improves performance.4,11 As an example, a monolithic polymer layer has been developed for PPEC by Nurok and co-workers.7 Separations of peptides and oligonucleotides were performed in 1−2 min on the 125 μm thick polymer monolith using a PPEC apparatus to control the separation temperature. Band Broadening. Separation efficiency in planar chromatography is described by plate number (N) and plate height (H). To evaluate the performance of the electrospun stationary phase, eq 2 was used to calculate N,2 where Zs is the distance ⎛ Z ⎞2 N = 16⎜ s ⎟ ⎝ wb ⎠

Figure 4. Plate height values versus migration distance: (A) R101 (orange circles), R590 (green triangles), and R610 (purple squares); (B) KR (dark blue diamonds) and SR (light blue circles). Conditions: 25:25:50 ACN/2-PrOH/25 mM citrate buffer, pH 5.6 (v/v/v), run at 1 kV for 60, 75, 90, and 120 s for each distance value.

(2)

but rather a plateau value of Hmin, suggesting that longitudinal diffusion does not contribute greatly to H. Under the conditions used, the lowest H value (11.3 μm) was observed for the analyte with the longest migration distance, R590, as both υeo and υep are toward the cathode for this positively charged analyte. This H is comparable to Hmin values (10 μm) reported so far in much more sophisticated PPEC systems using HPTLC plates,6 in which Hmin approaches 2dp, where dp is the average diameter of the stationary phase particle.4,6,16,42 These data suggest that even lower plate heights are possible at longer distances using electrospun nanofiber plates,10 but with the current unpressurized PEC setup, longer separation times (>3 min) suffered from drying effects, causing irreproducible separations. Further improvements in performance of electrospun stationary phases are expected when they are combined with the temperature control and heat dissipation capabilities of PPEC. It is anticipated that H values for these 400 nm nanofibers, once incorporated with PPEC, could approach 0.8 μm (or 2dp). Effect of Nanofiber Alignment. The effects of fiber alignment are discussed in the SI. Overall, randomly placed nanofiber plates displayed high efficiency and selectivity but provided the lowest migration rates; highly aligned nanofibers plates displayed low efficiency and selectivity but provided the fastest migration rates; and moderately aligned plates offered high selectivity and promoted fast migration rates while maintaining considerable efficiency. These data suggest that moderately aligned stationary phases are ideal for fast and simple separations, while randomly placed stationary phases are ideal for more complex mixtures which may require smaller spots. Therefore, the remaining separations were performed on randomly placed nanofiber plates.

traveled by the analyte from the sample origin and wb is the developed spot width (peak width at base). H, which is inversely related to N, describes the band broadening in the separation and can be calculated for planar chromatography using eq 3.2

H=

Zs N

(3)

The effect of analyte migration distance on plate height was investigated (Figure 4). H initially diminished with increasing distance and began to plateau at the longest distance examined. Although H theoretically does not change with migration distance for forced-flow separations,41 the variation in H detected here has also been observed in other PPEC systems.10,16 The total variance of the sample zone is composed of the combined variance of sample application of the plate, the variance associated with dispersion during analyte migration, and the variance related to the detection mode of the sample.8 The initial spot size makes a considerable contribution to the developed spot size, and the apparent efficiency of the layer is distorted for analytes with short migration distances.8,16 As such, large H values at short migration distances can be attributed to the substantial share of variance of sample application, which diminishes as migration distance is increased.4,8,16 In comparison, for capillary-driven chromatography, H typically increases at long distances past an observed Hmin as dispersion becomes controlled by longitudinal diffusion as the flow rate of the mobile phase slows considerably.2 This drawback is not applicable in PEC, since flow rate of the mobile phase is independent of distance.10 Therefore, there is not an increase after Hmin 10965

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

Effect of Stationary Phase Thickness. A main attraction of using electrospinning to generate stationary phases for PEC is that the layer thickness is readily tunable. Thickness is controlled simply by collecting nanofibers for various times.20,23,27,26 For electrospun PAN stationary phases, mat thickness linearly increases with electrospinning time up to a limiting time when thickness begins to plateau.20 At longer times, although thickness increases very little, the mass of the layer continues to increase.23 Therefore, longer relative electrospinning times may not have a significant effect on increasing thickness after the observed plateau, but there is a definite increase in the volume of stationary phase (Vs). Here, the impact of stationary phase thickness on analyte migration distance was investigated using stationary phases collected at different times. Plots of migration distances versus electrospinning times are given in Figure 5,

Figure 6. Plate height values on randomly placed nanofiber plates generated at different electrospinning times: 5 min, 12 μm thick (blue bars) or 20 min, 27 μm thick (red bars). PEC was performed for 1 min on the 5 min plate and for 1.5 min on the 20 min plate to achieve similar migration distances. Conditions: 25:25:50 ACN/2-PrOH/ 25 mM citrate buffer, pH 5.6 (v/v/v), mobile phase run at 1 kV.

different stationary phase thicknesses when developed for the same time (1 min), Figure 6 demonstrates that 20 min stationary phases produced lower H values relative to thinner 5 min stationary phases when plates were developed to similar distances (developed for 1.5 and 1.0 min, respectively). These data suggest that thinner stationary phases spun at 5 or 10 min are ideal for fast and simple separations, while 20 min stationary phases are ideal for separations of more complex mixtures that require smaller spots. Comparison of UTLC and PEC. The contribution of electrophoresis to the separation mechanism allows for differences in selectivity relative to liquid chromatography. PEC was anticipated to offer diverse selectivity compared to TLC using the same stationary phase. Since high-quality separation of the same laser dyes using PAN nanofiber stationary phases in UTLC has not been achieved,20,23,25 incorporating electrophoretic effects with PEC should offer distinctly different selectivity to aid in separating these analytes because they vary in charge. Using the Same Mobile Phase. To demonstrate that the selectivity observed in PEC was due to the addition of electrochromatographic effects rather than the difference in mobile phase, randomly placed nanofiber stationary phases were used in UTLC separations using the same mobile phase as in PEC (25:25:50 ACN/2-PrOH/25 mM citrate buffer, pH 5.6 (v/v/v)). Although these are not the optimum conditions for UTLC, this separation highlighted the differences in selectivity offered by the same stationary phase used in PEC mode. Additionally, differences in mobile phase velocity could be compared directly since the same mobile phase was used. To obtain a similar migration distance range, plates were developed over 15 mm for UTLC (>8 min), and plates were developed for 2 min for PEC. There was a wide difference in selectivity between the separation modes using the same mobile phase (SI, Table S-3). In UTLC, the retention order is R590 > PM and KR > R101, R610, and SR. Alternatively, using this mobile phase in PEC, negatively charged SR and KR were most retained, followed by PM, R610, and R101, and then positively charged R590 was least retained. SR and R590 reverse order of retention even though both separations utilized the same mobile phase and stationary phase. To compare analysis times between PEC and UTLC, separation times for the least retained analytes in each mode were compared over various distances (Table 2). The greater than 4-fold decrease in analysis time over this short distance underscores the advantage of using forced-flow mobile phases in PEC compared to capillary-flow in UTLC. As

Figure 5. Migration distance of laser dyes versus electrospinning time of the stationary phase. Analytes: KR (dark blue diamonds), PM (red squares), R101 (orange circles), R590 (green triangles), R610 (purple squares), and SR (light blue circkes). Conditions: potential applied for 1 min, 25:25:50 ACN/2-PrOH/25 mM citrate buffer, pH 5.6 (v/v/v), run at 1 kV.

using 25:25:50 ACN/2-PrOH/25 mM citrate buffer, pH 5.6 (v/v/v). Nanofibers were collected at 5, 10, and 20 min to generate stationary phase thicknesses of 12, 25, and 27 μm, respectively.20,23 Migration distances clearly decreased with increased electrospinning times as the thickness of the stationary phase increased. The 40−170% increase in analyte retention on stationary phases generated at 20 min was directly attributed to increased Vs available for analyte adsorption. Retention of analytes can be described by eq 4,1,2 where k is the k=K

Vs Vm

(4)

retention factor, K is the equilibrium constant, and Vm is the volume of mobile phase. Each stationary phase was composed of the same material; therefore, K remained constant. Vs was larger for stationary phases electrospun at longer times,23 and accordingly these layers produced an increase in k. Increase in k directly corresponded to shorter migration distances on stationary phases produced at longer times. Additionally, spot sizes also decreased by 20−115% with increased electrospinning time (SI, Figure S-2). Although spot sizes are expected to increase with development distance, thicker stationary phases consistently yielded smaller spots than 5 min stationary phases, even when developed to a similar distance (SI, Table S-2) using 1.5 min separation time for the 20 min plate. Consequently, although there was not a significant difference in H values for 10966

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

distance for E-UTLC based on previous studies.25,26 Separation time for UTLC using this mobile phase was 10 min; this was particularly fast compared to other capillary-driven separations utilizing 2-PrOH, and this fast time is characteristic for E-UTLC.2,20,23,25 However, comparatively, it took 2 min for analytes to migrate over 15 mm in PEC. For UTLC, KR and SR migrated to distances which have been noted to be close to the Hmin for E-UTLC (∼20 mm);25,26 H values of 33.9 and 38.2 μm were achieved, respectively (Table 3). These were

Table 2. Separation Times Observed in PEC and UTLC for the Least Retained Analytea separation time (min) migration distance (mm)b,c

PECb

UTLCc

6.9 7.7 9.6 15.7

1.00 1.25 1.50 2.00

1.60 2.97 3.13 8.30

a Conditions: 25:25:50 ACN/2-PrOH/25 mM citrate buffer, pH 5.6 (v/v/v), mobile phase on randomly-placed nanofiber stationary phases electrospun for 20 min. bMigration distance of the least retained analyte in PEC (R590). cSolvent front migration distance, Zf, as well as migration distance of the least retained analyte in UTLC (SR).

Table 3. Plate Height Values (%RSD) of Laser Dyes in UTLC and PEC Using Optimum Mobile Phasesa plate height (μm) laser dye

discussed earlier, mobile phase velocity in PEC does not diminish over time when the effects of Joule heating are minimized; rather, the υeo is independent of distance and is expressed according to the Helmboltz−Smoluchowski equation (eq 5), υeo =

ε0εrζE η

KR R101 R590 R610 SR

κ 2Zf

Zs (Zf − Z0)

PEC 36.4 21.5 11.3 48.5 85.3

(12) (19) (35) (35) (25)

UTLC was performed over 25 mm using 90:10 2-PrOH/MeOH (v/v) in 10 min, and PEC was performed for 2 min using 25:25:50 ACN/2-PrOH/25 mM citrate buffer, pH 5.6 (v/v/v). Separations utilized randomly placed nanofiber plates electrospun for 20 min.

typical for this distance using randomly placed PAN nanofiber plates.25 For PEC, R590 and R101 migrated to distances which were noted to be close to the Hmin described in Band Broadening (∼15 mm); H values of 11.3 and 21.5 μm were achieved, respectively. These were 2−3 times lower than what was observed in UTLC. The lower H values of PEC reiterate the benefit of using a forced-flow mobile phase that does not diminish over increasing distances. Additionally, zone capacity, or separation number (SN), was calculated. SN describes the number of analytes which can be separated on a given plate.2 SN can be experimentally determined using eq 8, by plotting

(6)

front migration distance. If the effects of Joule heat were minimized by incorporating electrospun plates with a PPEC device, still longer separation times and development distances could be used, and the advantage of linear mobile phase velocity in PEC over UTLC is only expected to be further enhanced. Using Optimum Mobile Phases. The performance of the stationary phase operated in PEC was compared to that of UTLC using the optimum mobile phases for each. Randomly placed nanofiber plates were used to separate five laser dyes in UTLC over 25 mm using 90:10 2-PrOH/MeOH (v/v)23,25 and in PEC over 2 min using 25:25:50 ACN/2-PrOH/25 mM buffer, pH 5.6 (v/v/v). For UTLC, selectivity of analytes was evaluated by calculating the retardation factor (Rf) of each analyte using eq 7,2 where (Zf − Z0) is the total separation Rf =

(11) (30) (28) (13) (30)

a

(5)

where ζ is the zeta potential, E is the electric field strength, and η is the mobile phase viscosity. Conversely, capillary-driven mobile phase velocity in TLC, υtlc, is not constant, and its value diminishes with increasing separation distance according to eq 6,2,4,5 where κ is the velocity constant and Zf is the solvent υtlc =

UTLC 36.8 149.3 131.3 80.7 38.2

SN =

(Zf − Z0) −1 (wH(start) + wH(front))

(8)

the spot widths at half height and extrapolating the values for analytes at the sample origin and end of the separation distance (wH(start) and wH(front), respectively).43SN = 7 was calculated for UTLC using the 25 mm separation distance, and SN = 11−12 was calculated for PEC in this same distance. The decreased H observed in PEC contributed to the enhanced SN relative to UTLC. It is also important to note that neither UTLC nor PEC could resolve all dyes in one dimension; however, combining these modes in a two-dimensional separation would be ideal, as they offer distinctly different selectivities. 2D UTLC-PEC. 2D chromatography uses two orthogonal methods of chromatography in series. Frequently in 2D-TLC, analytes are separated on the same plate, but development is performed with different mobile phases in perpendicular directions. 2D chromatography results in increased SN values which are critical in the separation of complex mixtures; the significance is rooted in the selectivity difference of the two dimensions. If the retention mechanisms are truly orthogonal, SN of the 2D separation (SN2D) is expected to be SN1 × SN2, where SN1 and SN2 correspond to the first and second dimensions, respectively.44 Typically, analytes are spotted at the bottom corner of a plate and developed in the first dimension. After drying, the same plate is turned 90° for development in the

(7)

distance. With this mobile phase, R590 and R101 were the most retained analytes, followed by R610, and KR and SR were least retained (Rf values were 0.02, 0.04, 0.16, 0.81, and 0.85, respectively). Retention order in UTLC was distinctly different from the retention order in PEC (see Using the Same Mobile Phase), and it agreed well with previous data using similar UTLC conditions.23,25 Two groups of analytes, those of highest and lowest retention, could not be resolved using this mobile phase in UTLC alone, which reiterates the need for a separation offering different selectivity. The separation distance used for UTLC (25 mm) was relatively short compared to that of conventional TLC; however, this was within the optimum 10967

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

dimension. SN = 7 was calculated for UTLC (SN1,UTLC), and SN = 10−11 was calculated for PEC (SN2,PEC); the combined capacity (SN2D) was in the range of 70−77. This was achieved in only 11 min (10 and 1 min for UTLC and PEC, respectively). Chomicki et al.46 have reported a greater capacity in the range of 150−200 in 2D HPTLC-PPEC; however, the plate was more than 6 times larger and the separation required 24 min. The high efficiency of E-UTLC plates allowed separations over shorter distances and thus shorter times. These separations not only confirmed the attractive selectivity difference available in PEC but also highlighted the advantage of utilizing E-UTLC plates rather than HPTLC plates in 2D planar chromatography separations.

second dimension using a different mobile phase for diverse selectivity. The combination of electrospun UTLC with PEC offers a promising approach to achieve a 2D planar chromatography separation on the same stationary phase but with different selectivity; however, few instances of TLC or HPTLC coupled with PEC have been reported,45−48 and this is the first time UTLC has been coupled with PEC. The separation power of a complex sample mixture using electrophoretic effects of PEC in combination with high efficiency of UTLC on electrospun stationary phases was examined. Randomly placed nanofiber plates (40 × 40 mm) were utilized to separate dyes using 2D UTLC-PEC (SI, Figure S-2B). Dyes were first separated by UTLC over 25 mm using 90:10 2-PrOH/MeOH.23,25 Once dried, the plate was developed by PEC for 1 min using the optimum mobile phase. Initial separations of six dyes resulted in tailed UTLC separations, as this was likely over the sample capacity of the phase.20 PM was removed from the mixture by lowering the concentration from 1.3 × 10−3 to 3 × 10−4 M, and separations of remaining dyes did not overload the plate. Figure 7 (SI, Figure S-6 provides a higher contrast

■

CONCLUSION Electrospun nanofiber stationary phases were successfully applied to PEC for the first time. Separations were achieved on a PAN nanofiber UTLC plate in 1−2 min using a ternary mobile phase. Buffer concentration was found to affect the rate of EOF, and an appropriate concentration was noted to balance separation speed with the effects of Joule heating. This system displayed good flow rate linearity between 60 and 90 s. Band broadening was characterized as a function of distance, and H as low as 11.3 μm was observed. Nanofiber alignment significantly increased migration rates, but lower H values were observed relative to those of randomly placed nanofibers. Another advantage of this thin layer is that the easily tunable stationary phase thickness can be tailored to a given separation. Compared to UTLC, PEC offered unique selectivity and decreased analysis times. Additionally, SN in the range of 70−77 was achieved in a rapid 2D UTLC-PEC separation. Electrospun phases showed considerable promise in PEC using this relatively simple setup; however, the true potential of the layer will be revealed once it is combined with PPEC.

■

Figure 7. Digital image of the electrospun stationary phase after a 2D UTLC-PEC separation of five laser dyes. The analyte origin was at the bottom corner. UTLC was performed first using 90:10 2-PrOH/ MeOH (v/v) and over 25 mm. PEC was performed second (perpendicular to UTLC) for 1 min using 25:25:50 ACN/2-PrOH/ 25 mM citrate buffer, pH 5.6 (v/v/v), at 1 kV. The colors in the image have been inverted and contrast enhanced for clarity. Analytes: (1) SR, (2) KR, (3) R610, (4) R101, and (5) R590.

ASSOCIATED CONTENT

* Supporting Information S

A more detailed description of the preparation of the electrospun stationary phases as well as specifics for UTLC development methods; Figures S-1−S-6 and Tables S-1−S-4 to support the data described within the text. This material is available free of charge via the Internet at http://pubs.acs.org.

■

image) shows an image of a plate after a 2D UTLC-PEC separation of five dyes. KR and SR were least retained in UTLC and were most retained in PEC; furthermore, R101 and R590 co-eluted in UTLC but were separated after PEC. This implied that the distinct selectivities were ideal for combination in 2D planar chromatography.46 The performance of PEC in the second dimension remained relatively high: H = 36.3 μm was observed for R590, the farthest migrating analyte, and H = 42.4 and 45.6 μm were observed for R610 and R101, respectively. These were comparable to the efficiency of using PEC in a single dimension (Figure 4A). Spot sizes after UTLC (first dimension) serve as starting spots for PEC (second dimension), and these were obviously larger than the starting spots obtained with the capillaries in the initial application. Due to increased spot sizes after UTLC, it would be reasonable to anticipate that the performance of the subsequent separation should be less efficient relative to the same separation used in one dimension. However, starting spots for PEC underwent a focusing effect during pre-wetting of the plate, as spots were wetted from either side.46 This effect was responsible for the H values observed, and it is incredibly advantageous with respect to using PEC in the second

AUTHOR INFORMATION

Corresponding Author

*Phone: +1-614-292-0733. Fax: +1-614-688-5402. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■ ■

ACKNOWLEDGMENTS The authors thank the National Science Foundation for funding this research under Grant NSF CHE-1012279. REFERENCES

(1) Poole, C. F. The Essence of Chromatography; Elsevier: Amsterdam/Boston, 2002. (2) Spangenberg, B.; Poole, C. F.; Weins, C. Quantitative Thin-Layer Chromatography: A Practical Survey; Springer-Verlag: Heidelberg, Germany, 2011. (3) Hauck, H. E.; Bund, O.; Fischer, M.; Schulz, M. J. Planar Chromatogr. 2001, 14, 234. (4) Dzido, T. H.; Plocharz, P. W.; Chomicki, A.; Halka-Grysinska, A.; Polak, B. J. Chromatogr. A 2011, 1218, 2636. 10968

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969

Analytical Chemistry

Article

(5) Dzido, T. H.; Plocharz, P. W.; Slazak, P.; Halka, A. Anal. Bioanal. Chem. 2008, 391, 2111. (6) Plocharz, P.; Klimek-Turek, A.; Dzido, T. H. J. Chromatogr. A 2010, 1217, 4868. (7) Woodward, S.; Urbanova, I.; Nurok, D.; Svec, F. Anal. Chem. 2010, 82, 3445. (8) Dzido, T. H.; Plocharz, P. W. J. Liq. Chromatogr. Relat. Technol. 2007, 30, 2651. (9) Nurok, D.; Koers, J. M.; Novotny, A. L.; Carmichael, M. A.; Kosiba, J. J.; Santini, R. E.; Hawkins, G. L.; Replogle, R. W. Anal. Chem. 2004, 76, 1690. (10) Dzido, T. H.; Plocharz, P. W.; Slazak, P. Anal. Chem. 2006, 78, 4713. (11) Chomicki, A.; Dzido, T. H.; Plocharz, P.; Polak, B. In Electrochromatography Methods: Planar Electrochromatography; Buszewski, B., Ed.; Springer-Verlag: Berlin, 2013; p 191. (12) Berezkin, V. G.; Balushkin, A. O.; Tyaglov, B. V.; Rozhilo, Y.; Malinovska, I. Dokl. Phys. Chem. 2004, 399, 287. (13) Dzido, T. H.; Majewski, R.; Polak, B.; Golkiewicz, W.; Soczewinski, E. J. Planar Chromatogr. 2003, 16, 176. (14) Patton, W. F.; Panchagnula, V.; Rockney, E.; Krull, I. S. J. Liq. Chromatogr. Relat. Technol. 2006, 29, 1177. (15) Dzido, T. H.; Mroz, J.; Jozwiak, G. W. J. Planar Chromatogr. 2004, 17, 404. (16) Novotny, A. L.; Nurok, D.; Replogle, R. W.; Hawkins, G. L.; Santini, R. E. Anal. Chem. 2006, 78, 2823. (17) Tate, P. A.; Dorsey, J. G. J. Chromatogr. A 2005, 1079, 317. (18) Tate, P. A.; Dorsey, J. G. J. Chromatogr. A 2006, 1103, 150. (19) InChromatics: The Ultimate Source in High-Throughput Separations. www.inchromatics.com (accessed January 16, 2014). (20) Clark, J. E.; Olesik, S. V. Anal. Chem. 2009, 81, 4121. (21) Andrady, A. L. Science and Technology of Polymer Nanofibers; John Wiley & Sons, Inc.: Hoboken, NJ, 2008. (22) Clark, J. E.; Olesik, S. V. J. Chromatogr. A 2010, 1217, 4655. (23) Beilke, M. C.; Zewe, J. W.; Olesik, S. V. Anal. Chim. Acta 2013, 761, 201. (24) Kampalanonwat, P.; Supaphol, P.; Morlock, G. E. J. Chromatogr. A 2013, 1299, 110. (25) Fang, X.; Olesik, S. V. Anal. Chim. Acta 2014, 830, 1. (26) Newsome, T. E.; Olesik, S. V. J. Chromatogr. A 2014, 1364, 261. (27) Lu, T.; Olesik, S. V. J. Chromatogr. B 2013, 912, 98. (28) Rojanarata, T.; Plianwong, S.; Su-uta, K.; Opanasopit, P.; Ngawhitunpat, T. Talanta 2013, 15, 208. (29) Miller, J. M. Chromatography: Concepts and Contrasts, 2nd ed.; Wiley: Hoboken, NJ, 2005. (30) Merouani, S.; Hamdaoui, O.; Saoudi, F.; Chiha, M. Chem. Eng. J. 2010, 158, 550. (31) Lahnstein, K.; Schmehl, T.; Rusch, U.; Rieger, M.; Seeger, W.; Gessler, T. Int. J. Pharm. 2008, 351, 158−164. (32) Gilliland, J. W.; Yokoyama, K.; Yip, W. T. Chem. Mater. 2004, 16, 3949−3954. (33) Subirats, X.; Roses, M.; Bosch, E. Sep. Purif. Rev. 2007, 36, 231. (34) Kazakevich, Y.; LoBrutto, R. HPLC for Pharmaceutical Scientists; John Wiley & Sons: Hoboken, NJ, 2007; pp 139−240. (35) Nurok, D.; Koers, J. M.; Nyman, D. A.; Liao, W. M. J. Planar Chromatogr. 2001, 14, 409. (36) Cahours, X.; Morin, P.; Dreux, M. J. Chromatogr. A 1998, 845, 203. (37) Nurok, D.; Koers, J. M.; Carmichael, M. A. J. Chromatogr. A 2003, 983, 247. (38) Polak, B.; Halka, A.; Dzido, T. H. J. Planar Chromatogr. 2008, 21, 33. (39) Polak, B.; Wojtanowski, K. K.; Slazak, P.; Dzido, T. H. Chromatographia 2011, 73, 339. (40) Kasnavia, T. N.; Vu, D.; Sabatini, D. A. Goundwater 1999, 37, 376. (41) Poole, C. F. J. Chromatogr. A 1999, 856, 399. (42) Plocharz, P. W.; Dzido, T. H.; Slazak, P.; Jozwiak, G. W.; Torbicz, A. J. Chromatogr. A 2007, 1170, 91.

(43) Hall, J. Z.; Taschuk, M. T.; Brett, M. J. J. Chromatogr. A 2012, 1266, 168. (44) Milz, B.; Klein, K. F.; Spangenberg, B. J. Planar Chromatogr. 2013, 25, 493. (45) Panchagnula, V.; Mikulskis, A.; Songa, L.; Wang, Y.; Wang, M.; Knubovets, T.; Scrivener, E.; Golenko, E.; Krull, I. S.; Schulz, M.; Hauck, H. E.; Patton, W. F. J. Chromatogr. A 2007, 1155, 112. (46) Chomicki, A.; Slazak, P.; Dzido, T. H. Electrophoresis 2009, 30, 3718. (47) Chomicki, A.; Kloc, K.; Dzido, T. H. J. Planar Chromatogr. 2011, 24, 6. (48) Stevenson, P. R.; Dunlap, B. E.; Powell, P. S.; Petersen, B. V.; Hatch, C. J.; Chan, H.; Still, G. I.; Fulton, M. T.; McKell, J. S.; Collins, D. C. Anal. Bioanal. Chem. 2013, 405, 3085.

10969

dx.doi.org/10.1021/ac503568a | Anal. Chem. 2014, 86, 10961−10969