Inline Coupling of Electrokinetic Preconcentration Method to Taylor

Jan 23, 2018 - (42) Indeed, as recommended by Pecora, in all instances, Dh,DLS should be calculated from the self-diffusion coefficient at infinite di...
2 downloads 10 Views 2MB Size
Download PDF
Article pubs.acs.org/ac

Cite This: Anal. Chem. XXXX, XXX, XXX−XXX

Inline Coupling of Electrokinetic Preconcentration Method to Taylor Dispersion Analysis for Size-Based Characterization of Low-UVAbsorbing Nanoparticles Farid Oukacine,*,† Annabelle Gèze,† Luc Choisnard,† Jean-Luc Putaux,‡ Jean-Paul Stahl,§ and Eric Peyrin† †

Univ. Grenoble Alpes, DPM, CNRS UMR 5063, F-38041 Grenoble, France Univ. Grenoble Alpes, CNRS, CERMAV, F-38000 Grenoble, France § Infectiologie, Univ. et CHU Grenoble Alpes, 38700 La Tronche, France ‡

S Supporting Information *

ABSTRACT: The inline coupling of the field-amplified sample injection (FASI) to Taylor dispersion analysis (TDA) was used to characterize low-UV absorbing carboxylated silica nanoparticles (cNPs). The hydrodynamic diameters (Dh) were measured by using a commercial capillary electrophoresis instrument. The proposed methodology did not require any complicated instruments or chromophoric dye to increase the detection sensitivity. A practical method based on a halfGaussian fitting was proposed for the data processing. The results obtained by this method were compared with those derived from dynamic light scattering (DLS) and transmission electron microscopy (TEM) analyses. From these results, it appeared that the size derived by TDA is in excellent agreement with those measured by DLS and TEM, as demonstrated by stable nanoparticles with narrow size distributions. Intermediate precision relative standard deviations less than 5% were obtained by FASI-TDA. The effect of the FASI-induced cNP peak dispersion on the reliability of the results was discussed in detail.

N

have been published over the last years.19−22 The advantage of combining complementary methods is clearly highlighted. Another method allowing the determination of the diffusion coefficients, and hence the hydrodynamic diameters (Dh) of NPs is the Taylor dispersion analysis (TDA). This method, described by Taylor in 1953,23 is based on a dispersion of a solute plug in a capillary under laminar Poiseuille flow. After injection in the capillary of a solute dissolved in the eluent, a pressure is applied at the end of the capillary. Solutes in the band move with different velocity depending on their position in the capillary because the velocity profile is a parabolic function of the radius. The combination of the dispersive velocity profile with the molecular diffusion that redistributes solutes in the capillary cross section leads to a specific mechanism of dispersion described by the Taylor−Aris equation.24 The analysis can be carried out either at a single detection point or at two spatially separated detection points.25 TDA has recently gained interest. Indeed, an automated biophysical characterization tool using TDA is commercially available (Malvern Panalytical Viscosizer TD). TDA has been used for a wide variety of structures including peptides and proteins,26−28 small molecules,29 macromolecules,30 and nano-

anoparticles (NPs) are of great scientific interest because they constitute a bridge between bulk materials and molecular structures.1 During the last decades, NPs have been intensively used in many areas such as electronics, cosmetics, energy, chemistry, biology, medicine, and food.2−4 More recently, the environmental impact and toxicity of nanomaterials have raised the attention of the researchers and social community. Indeed, due to their small size, NPs exhibit a high level of cell penetration through different barriers.5 Several adverse effects of nanomaterials on human health have already been reported.6 Therefore, from a regulatory point of view, a rigorous and practical method for enabling reliable characterization of nanomaterials is essential.7 A variety of techniques are available to characterize the size and size distribution of NPs. These include transmission electron microscopy (TEM),8 atomic force microscopy (AFM),9 scanning electron microscopy (SEM),10 dynamic light scattering (DLS), 11 analytical ultracentrifugation (AUC),12 size-exclusion chromatography (SEC),13 asymmetrical flow field-flow fractionation (AF4),14 hydrodynamic chromatography (HDC),15 X-ray diffraction (XRD),16 and small-angle X-ray scattering (SAXS).17 All these analytical methods present advantages as well as limitations, and there is no available technique allowing the analysis of all kinds of nanomaterials.18 Several publications describing the advantages and the limitations of the commonly used analytical methods © XXXX American Chemical Society

Received: August 17, 2017 Accepted: January 23, 2018 Published: January 23, 2018 A

DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX

Article

Analytical Chemistry particles.31−33 TDA was also used for the analysis of noncovalent binding interactions.34,35 However, like all the other methods, TDA has its own limitations. For example, TDA can easily be disturbed by the presence of chromophoric compounds (e.g., UV-absorbing solvent, impurity) in the sample.24 Interactions between solutes and capillary walls can be detrimental for the results, and the capillary wall interactions must be minimized.36 Moreover, as is the case for the DLS, TDA gives the size of one diffusing ensemble. For the mixture of solutes, or polydisperse samples, TDA leads to a harmonic weight-average diffusion coefficient.37 Recently, methodologies have been developed for the analysis of sample polydispersity38,39 and for the analytical mitigation of solute−capillary wall interactions.36 Moreover, by applying an initial electrophoretic separation step prior to the application of the pressure, it is possible to perform individual diffusion coefficient measurements in a sample mixture.28,40 Despite these achievements, when the UV detection is used, TDA cannot allow the analysis of low-UV-absorbing particles and/or molecules. More recently, a backscattering interferometer was coupled to TDA for the analysis of non-UVabsorbing polymers.41 However, this method requires the modification of the CE cartridge, the use of CCD camera, He− Ne laser as well as a Peltier thermoelectric cooler. In this work, the inline coupling of electrokinetic preconcentration method to TDA was assessed for characterizing low-UV-absorbing carboxylated nanoparticles (cNPs). The proposed methodology did not require any complicated instruments or chromophoric dye to increase the detection sensitivity. Three batches of cNPs with nominal size of 10, 30, and 50 nm were analyzed. A practical method allowing the data processing was proposed and the obtained results were compared to those derived from DLS and TEM.

by using the same dispersant. In our experiments, the size of the cNPs was measured at different concentrations (Supporting Information Figure S-1). Subsequently, extrapolations of the apparent measured size to zero concentration were carried out to obtain concentration-independent sizes.42 As described in Table 1, the results obtained for cNP-30 are in good agreement with those reported by Micromod Corp. However, the cNP−50 diameter was lower than that provided by the manufacturer. Indeed, the silica nanoparticles have inherent dissolution characteristics within aqueous systems43,44 which could explain this discrepancy. For cNP−10, we observed a difference of 2.3 nm between our results and those reported by the manufacturer. However, these NPs are highly polydisperse in pure water ( 0.272) and DLS is not suitable when dealing with such samples.20 Capillary Electrophoresis. Capillary electrophoresis (CE) experiments were carried out with a 3D-CE instrument (Agilent Technologies, Waldbronn, Germany) equipped with a diode array detector. Bare-fused silica capillaries were purchased from Photon lines (Saint-Germain-en-Laye, France). Poly(vinyl alcohol) (PVA) coated capillaries were purchased from Interchim (Montluçon, France). New bare-fused silica capillaries were conditioned by performing the following washes: 1 M NaOH for 30 min and water for 10 min. The washing process between runs was performed with 1 M NaOH (10 min), H2O (2 min), and electrolyte (5 min). For the experiments performed in PVA capillaries, the washing process between runs was performed with water (2 min) and electrolyte (5 min) at 1 bar. At the end of each day, the PVA capillaries were flushed with water for 20 min and stored at 4 °C. The BGE buffer consisted of Tris/acetic acid 35.9/21.2 mM (pH 7.8). Dynamic Light-Scattering Measurements. DLS measurements were carried out with a Malvern Zetasizer Nano ZS ZEN 3500 (laser wavelength 532 nm) in general purpose mode (with normal resolution). The scattering was monitored at a fixed angle of 173° in backward scattering mode. The equilibration time before each triplicate was fixed at 120 s. The water viscosity (0.89 mPa s at 25 °C) was used as sample viscosity. cNPs were analyzed in BGE buffer. Transmission Electron Microscopy. For TEM observations, droplets of suspensions of cNP‑10 (0.01 mg mL−1), cNP‑30 and cNP‑50 (0.1 mg mL−1) nanoparticles diluted in BGE were deposited onto glow-discharged carbon-coated microscopy grids. The excess liquid was blotted and the samples were observed with a Philips CM200 microscope (FEI, Hillsboro, U.S.A.) operating at 200 kV. The images were recorded with a TemCam F216 digital camera (TVIPS, GmbH, Germany). The nanoparticle diameter was measured from populations of 300 particles using the ImageJ software and the number-average mean diameter (DN,TEM) was determined by using eq 1. The weight-average (DW,TEM) and Z-average (DZ,TEM) mean diameters were calculated from the series of diameters by using eqs 2 and 3 respectively:45

■

EXPERIMENTAL SECTION Chemicals. Tris(hydroxymethyl)aminomethane (Tris) and 2-(N-morpholino)ethanesulfonic acid (MES) were purchased from-sigma-Aldrich (Saint-Quentin, France). Acetic acid and NaOH were provided by Carlo Erba (Val de Reuil, France). H3PO4 was from Fisher Scientific (Illkirch-Graffenstaden, France). Na2CO3 was from Fluka Chemie AG (Buchs, Switzerland). Water was purified using a Purite Still Plus system (Thame, U.K.). Three batches of cNPs: cNP-10, cNP-30, and cNP-50 at 25 mg mL−1 were purchased from micromod Partikeltechnologie GmbH (Rostock, Germany). A propyl spacer was used by the manufacturer for the carboxylic functionalization of the silica NPs. The sizes of the freshly synthesized cNPs as reported by the manufacturer are provided in Table 1. Pure water was used as dispersant. At first, in order to evaluate the accuracy of our DLS data, we analyzed the cNPs Table 1. Size of the cNPs Measured by DLS by Using Water as Dispersant cNP‑10 cNP‑30 cNP‑50

product codes

Dh,DLS (nm)a

b

Dh,DLS (nm)c

PdIc

43-02-101 43-02-301 43-02-501

11.3 30.4 45.0

0.272 0.069 0.069

9 31 53

0.350 0.050 0.080

D N,TEM =

∑i ND i i ∑i Ni

(1)

a

Hydrodynamic diameters calculated from the extrapolations of the apparent measured sizes to zero concentrations. bAverage PdIs recorded during the experiments (Supporting Information Figure S1). cSizes of the freshly synthesized cNPs given by the manufacturer.

D W,TEM = B

4 ∑i ND i i 3 ∑i ND i i

(2) DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX

Article

Analytical Chemistry

Figure 1. Size distribution of the cNPs determined by DLS with their corresponding TEM images. (A) Dh distributions recorded by DLS at multiple concentrations of NPs. (B) Plot of the Dh,DLS vs NPs concentration where the y-axis intercepts provide concentration-independent Dh. (C) Corresponding TEM images of the cNPs. BGE was used as dispersant. Other experimental conditions are described in the Experimental Section.

DZ,TEM =

6 ∑i ND i i 5 ∑i ND i i

zero concentration were carried out to obtain concentrationindependent sizes (Figure 1B).42 Indeed, as recommended by Pecora, in all instances, Dh,DLS should be calculated from the self-diffusion coefficient at infinite dilution.46 The Z-average mean diameter of the cNP‑10, cNP‑30, and cNP‑50 are 16.9 ± 0.8, 30.7 ± 1.5, and 42.5 ± 2.1 nm, respectively. Because of various systematic errors, we used 5% uncertainty for the Dh,DLS.47−49 TEM images (Figure 1C) show that both cNP‑30 and cNP‑50 nanoparticles are fairly spheroidal and uniform in size, in good agreement with the low PdI values obtained by DLS. However, as compared with cNP‑50, cNP‑30 particles are slightly more polygonal in shape. The DZ,TEM values of cNP‑50 and cNP‑30 (35.9 and 25.2 nm, respectively) are lower than those calculated by DLS. Indeed, DLS measures the hydrodynamic diameter of the particles which includes other molecules and ions that surround the particle core and move together in solution.50 As reported by Shaw, the hydrodynamic diameter measured by DLS depends not only on the size of the particle “core”, but also on any surface structure, as well as the type and concentration of any ions in the medium.51 The probable presence of the propyl shells on the surface of the particle may also affect the translational friction coefficient and change the apparent size of the particle.51 Moreover, small molecular size organic compounds are electron-transparent, and therefore, they cannot be seen in the TEM micrograph.22,52 Differences

(3)

where Ni is the number of particles of the diameters Di. This allows for a better comparison with the weight-average and Zaverage mean diameters measured by TDA37 and DLS, respectively.

■

RESULTS AND DISCUSSION Size Characterization of cNPs by DLS and TEM. BGE is required in FASI-TDA experiments to provide the transport of the electric current and the separation of the analytes. Then, for a better comparison with the results derived from FASI-TDA, cNPs were characterized by DLS and by TEM in the same dispersion medium (Tris/acetic acid 35.9/21.2 mM, pH 7.8). Figure 1A shows the size distribution profiles of the cNPs at multiple concentrations. Polydispersity indexes (PdIs) obtained in these experimental conditions were lower than 0.100 suggesting a narrow particle size distribution for all cNPs. It is interesting to notice that the PdI of cNP-10 ( 0.100) measured in these conditions is lower than that obtained in pure water ( 0.272). This suggests that the cNP-10 aggregate, in a reversible way, in water. The size of the cNPs was measured by DLS at different concentrations (Figure 1A). Subsequently, extrapolations of the apparent measured size to C

DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX

Article

Analytical Chemistry

Figure 2. Taylorgrams of three individual carboxylated silica nanoparticles of different sizes at (A) 195 nm, (B) 300 nm. Experimental conditions: Poly(vinyl alcohol) (PVA) coated capillary 65 cm (56.5 cm to the detector) × 50 μm i.d. T = 25 °C. Hydrodynamic injection: 50 mbar, 5 s. Mobilizing pressure: 50 mbar. cNPs concentration: 10 mg mL−1 diluted in the BGE. Other conditions are described in the Experimental Section.

Figure 3. Inline coupling of the FASI to TDA for the cNPs characterization. Experimental conditions: Poly(vinyl alcohol) (PVA) coated capillary 65 cm (56.5 cm to the detector) × 50 μm i.d. T = 25 °C. Sample: cNP‑30 at 10 mg mL−1 in 10-times diluted BGE. 1st step: Electrokinetic injection: −10 kV, 60 s. Applied voltage: −20 kV during 9.7 min. 2nd step: −50 mbar for 8 min with BGE at the inlet and at the outlet of the capillary. After that, the mobilization pressure was reversed (+50 mbar) to allow the double detection of the cNPs. Other conditions are described in the experimental section. Abbreviations: Applied voltage (V), mobilization pressure (ΔP).

were analyzed by conventional TDA. Figure 2A described the Taylorgrams recorded at 195 nm. cNPs were analyzed in poly(vinyl alcohol) (PVA) coated capillary at a concentration of 10 mg mL−1. As described in Figure 2A, the profiles of the Taylorgrams are the sum of two distinct distributions, as typically observed for binary mixtures with analytes of largely different sizes.37 The elution profiles of the peaks marked with asterisks are narrower than those corresponding to the cNPs. This demonstrates that the cNPs samples are not pure and contain other small molecules/ions (e.g., synthetic chemical residues, salts, etc.). Further analysis of the cNPs by capillary zone electrophoresis (CZE) demonstrated that these species are mostly positively charged and neutral (Supporting Information Figure S-2). The presence of these compounds and the low-UV absorbance of the cNPs associated with the baseline deformation hinder the exploitation of the data (Supplementary Figure 2). By using a selective wavelength (300 nm, see Figure 2B), small molecules are not detected. However, the cNP signals are dramatically reduced. Inline Coupling of the Field Amplified Sample Injection to Taylor Dispersion Analysis. To further improve the sensitivity of the detection, the field-amplified sample injection (FASI) was coupled to TDA. This type of stacking is based on differences in the electric field between the sample and the electrolyte. Moreover, when FASI is performed under conditions in which there is no electroosmotic flow, it is

between hydrodynamic radius and TEM primary particle radius are then expected. For cNP-10, TEM images show individual primary particles as well as more irregular aggregates of 2−4 units. However, DZ,TEM (16.6 nm) calculated from the distribution in the images is close from that derived by DLS (16.9 ± 0.8 nm). At first sight, for the reasons mentioned above, it is reasonable to assume that the larger particles do not exist in suspension and are drying artifacts.53 However, this comparison must be treated with some degree of caution because cNP-10 exhibits nonspherical morphologies. It should be noted that, in DLS, the hydrodynamic diameter measured for the particle is that of a hypothetic hard sphere that moves at the same velocity in solution as the particle in question. For this reason, the sizes obtained by DLS may not be representative for nonspherical morphologies.54 Taylor Dispersion Analysis of Low-UV-Absorbing cNPs. As discussed in the introduction, TDA is based on the dispersion of a solute plug in capillary under a laminar Poiseuille flow. After injection in the capillary of a solute dissolved in the eluent, a pressure is applied at the end of the capillary. The temporal variance of the elution peak can be theoretically deduced either by fitting the Taylorgrams by a Gaussian,35 by integration of the elution profile55 or by calculating the profile moments.28 However, in the case of the cNPs, the obtained Taylorgrams are not exploitable. For example, in the initial experiments, the three batches of cNPs D

DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX

Article

Analytical Chemistry

Figure 4. Taylorgrams of cNP‑30, cNP‑50, and cNP‑10 obtained with dual-UV detection system by inline coupling of FASI to TDA (A). Graphical illustration of the cNPs hydrodynamic diameters obtained with different sets of PVA capillaries (B). Experimental conditions are described in Figure 3 and in the Experimental Section. σI is the intermediate precision standard deviation related to the overall averages obtained by TDA.

preparation in the laboratory is laborious. The data processing of such peaks requires an alternative approach to the commonly used Gaussian fitting35 or integration method that consists in determining the temporal variance by integration of the whole signal.55 In this work, a practical method based on a halfGaussian fitting is proposed. This alternative approach does not require experimental mitigation of the artifact. The fits of the Taylorgrams were realized with Microcal Origin 6.0 by using the following GAUSS Equation (eq 4):

possible to perform long electrokinetic injection (EKI) while keeping the physical volume of the sample matrix entering the capillary very low.56,57 A 100−1000-fold sensitivity improvement is generally reached as compared with a hydrodynamic injection. More recently, the EKI of carboxylated magnetite nanoparticles and their online preconcentration was reported by Baron et al.58 The dual-UV detection system used in this work is similar to that reported by Leclercq et al.40 The experiments were performed on a commercial capillary electrophoresis apparatus and did not require an external UV detector.28 In the first step of the method (Figure 3A), cNP‑30 at 10 mg mL−1 suspended in 10-times diluted BGE were injected in the capillary by EKI. As described previously, the aim of this dilution is to increase the resistivity of the cNPs as compared to the electrolyte. A voltage (V) was next applied until the cNP plug traveled a distance corresponding to ∼97.7% of the total capillary length. Then, the voltage was turned off. The stop time can easily be calculated by using the time required for the plug to reach the detector. A typical electropherogram of the cNP‑30 recorded at 300 nm is depicted in Figure 3A. As expected, peak intensity obtained in FASI mode is much higher than that obtained by using conventional HDI. Moreover, the peak profile was greatly improved. It should be noted that the FASI of cNP-30 at a concentration of 2.5 mg mL−1, in these experimental conditions, induces the breakdown of the current. This is due to the very low sample conductivity in these conditions and to a long injection time (60 s) which both cause the electrophoretic process instability. However, for this range of concentrations (∼2.5 mg mL−1), the EKI can easily be performed by suspending the cNP‑30 in 5times diluted electrolyte (Supporting Information Figure S-3). In the second step of the method (Figure 3B), a negative mobilization pressure (ΔP) was applied to move the NPs toward the capillary inlet. After 8 min, the mobilization pressure was reversed to allow the double detection of the NPs and a differential measurement of the peak dispersion. Data Processing of the Taylorgrams. As depicted in Figure 3B, an artifact (marked with an asterisk) is present at the left and the right sides of the cNP bands. This artifact is related to the baseline falling down immediately after the application of the negative mobilization pressure. To pull the cNPs bands away from this artifact, the distance between the detection cell and the capillary outlet should be increased and the capillary cartridge should be modified in the same way than that reported by Tohala et al.59 However, the PVA-coated capillaries are available in the market in standard dimensions and their

(x − xc) A −2 w2 y = y0 + e w π /2

2

(4)

where A is the area, w is defined as 2 times the standard deviation of the Gaussian fit (2σ), xc is the location parameters (average migration time of the peak), and y0 is the baseline offset. The average migration time of the peak was used to constrain the fitting algorithm. This parameter was estimated from the position of the cNP peak apex. The detailed protocol of the data processing is described in the supplementary Figure 4. Trace (a) of Figure 4A, displays an example of the obtained results after the data processing of the Figure 3B Taylorgrams. As described in this figure, only the right and the left sides of the cNP‑30 bands were taken into account for the fits (blue dashed areas of the Taylorgrams). The data processing were performed on the Taylorgrams recorded at 300 nm for all the cNPs (Supplementary Figure 4). Fit parameter values are shown on the graph. It should be noted that the parameter w calculated by Origin is not that of the truncated Gaussian, but it corresponds to that of the whole curve. Subsequently, the diffusion coefficient (D) can be calculated by using eq 5.28,40,55 D=

(

R C2 t 2 − t1 − 24(σ22

−

t ramp

2 2 σ1 )

) (5)

where Rc is the capillary radius and tramp is the time during which the pressure increases linearly from −50 to +50 mbar in the second step of the method. This value is an apparatusrelated constant which is equal to 7.2 s. σ21 and σ22 are the peak variances at the first and at the second detection. t1 and t2 are the experimentally observed detection times. Hydrodynamic diameters are determined using eq 6: Dh = E

kBT 3πηD

(6) DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX

Article

Analytical Chemistry Table 2. Size of the Carboxylated Silica Nanoparticles Measured by TDA, DLS, and TEM FASI-TDAa

DLSb

TEMc

sample

n=12 (nm)

RSDI,n=12 (%)

DLS (nm)

PdI

DN,TEM (nm) [std nm]

DW,TEM (nm)

DZ,TEM (nm)

cNP‑10 cNP‑30 cNP‑50

15.6 ± 4.4 29.4 ± 2.4 37.7 ± 2.7

14.24 4.13 3.60

16.9 ± 0.8 30.7 ± 1.5 42.5 ± 2.1

0.100 0.048 0.060

12.3 [2.9] 21.2 [4.0] 31.7 [5.1]

14.6 23.5 34.2

16.6 25.2 35.9

Overall average hydrodynamic diameters (±2 × intermediate precision standard deviations) and intermediate precision relative standard deviation obtained by FASI-TDA. The experimental conditions for the FASI-TDA experiments are described in Figure 3. bHydrodynamic diameters and PdIs of the cNPs were measured with Zetasizer in BGE (pH 7.8). Because of various systematic errors, we used 5% uncertainty for the Dh,DLS. cThe experimental conditions of the TEM experiments are described in the Experimental Section. a

Figure 5. Taylorgrams of cNP‑30, cNP‑50, and cNP‑10 obtained with dual-UV detection system by inline coupling of EKI to TDA (A). Graphical illustration of the cNP hydrodynamic diameters obtained in different dilution media (B). NP samples are suspended in (A) BGE, (B) as described in the figure. Other experimental conditions are depicted in Figure 3.

where kB is the Boltzmann constant, T is the absolute temperature, and η is the viscosity of the eluent. Because of the low concentration of the BGE, the water viscosity was used as approximate for the calculation (η = 0.896 mPa s at 25 °C).60 It should be noted that the eq 5 is valid when the following equations are fulfilled:

(

D t 2 − t1 − R C2 R C3ΔP > 69 8ηLD

t ramp 2

intensity of the scattered light is proportional to the sixth power of the radius.21 However, in both cases, the measured hydrodynamic diameter includes the organic shell as well as the hydration layer present at the surface of the NPs. Therefore, in the case of monodisperse samples, close experimental values should be obtained by both methods. TEM images provide the size of the inorganic core of the NPs. Despite the conversion of the number mean diameter (DN,TEM) into weight-average mean diameter (DW,TEM), the measured size by TEM should be lower than that recorded by TDA and DLS. This was verified for the cNP‑30 and cNP‑50 (Figure 4B). Indeed, the Dh,DLS values of the cNP‑50 and cNP‑30 (42.5 ± 2.1 and 30.7 ± 1.5 nm) are equal to or slightly higher than those resulting from TDA (37.7 ± 2.7 and 29.4 ± 2.4 nm) while the DW,TEM values of the cNP‑50 and cNP‑30 are respectively 34.2 and 23.5 nm. Note that intermediate precision relative standard deviation (RSDI) less than 5% are obtained for both cNP‑30 and cNP‑50 by FASI-TDA (see Table 2). Moreover, R2 values for all the half-Gaussian fits related to these two NPs are in the range of 0.9971 to 0.9999. Principal Limitation of the Method. Contrary to what was obtained for cNP‑30 and for cNP‑50, the half-Gaussian shape of cNP‑10 was not observed at the first detection point (see trace (c) in Figure 4A). This hinders the accurate determination of the Dh,TDA by using the half-Gaussian fitting method and this leads to a broad spread of the size values (see Table 2). Indeed, although the n=12 measured (15.6 ± 4.4 nm) is the same than that derived from DLS (16.9 ± 0.8 nm), significantly larger RSDI (14.24%) was obtained. Then, the halfGaussian fitting method is not adapted for the data processing of such signal. This is the primary limitation of this method. Because the cNPs were introduced into the capillary by FASI, their concentrations in the sample zones are very high. As described in the first part of this manuscript, a low NP concentration induces the breakdown of the current during the

) ≥ 1.4 (7)

(8)

ΔP is the absolute value mobilization pressure, and L is the total capillary length. Note that the relation 8 is a linear combination of the Peclet number equation28 and Poiseuille’s law.61−63 Characterization of cNPs by the Inline Coupling of FASI to TDA. Typical Taylorgrams obtained for the cNP‑50 and for the cNP‑10 are also shown in traces (b) and (c) of Figure 4A, respectively. The experiments were performed in quadruplicate. Intermediate precision was assessed by using three different sets of PVA capillaries (PVA 1, PVA 2, PVA 3). Figure 4B displays the individual values (○), the average values per capillary (+) as well as the overall average values (in black lines) of the Dh,TDA measured for the three batches of the cNPs by FASI-TDA. The confidence intervals at 95% which are equal to ±2 × intermediate precision standard deviations (σI)64 are in dashed black lines in Figure 4B. For a better comparison, DZ,DLS (in red line) and DW,TEM (in purple line) are also depicted. As described in the previous section, DLS gives intensity-weighted harmonic mean size while TDA provides weight-average Dh.21,24 Dh,DLS is shifted to larger size particles because the F

DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX

Analytical Chemistry

■

experiment. It is consequently difficult to study the effect of the starting concentration on the peak broadening. Moreover, due to the preconcentration effect, the starting concentration does not reflect the real concentration of the NPs in the capillary. To evaluate the contribution of the FASI process to the finally recorded TDA signal, the same experiments were performed in absence of field amplification. The results are shown in Figure 5A. The only difference between Figure 5A and Figure 4A, is the dilution medium of the cNPs. In Figure 5A, the cNPs are diluted in BGE, whereas in Figure 4A, they are suspended in 10-times diluted BGE. As described in Figure 5A, in the absence of field amplification during the injection, the halfGaussian shapes are conserved in all cases. There is therefore no doubt about the fact that the deviation from the Gaussian curve, appearing at the apex of the cNP‑10 Taylorgram, is closely related to the high cNP‑10 concentration in the sample zone induced by the preconcentration step. However, high NP concentration in the sample zone can have several significant effects, all detrimental for the FASI-TDA analysis. For example, high NP concentration can exacerbate field inhomogeneities and generates distorted peak shapes caused by mismatched conductivity between the running buffer and the sample zone.65 At higher concentrations, particle interactions can also modify the free diffusion of particles and this can lead to nonspecific aggregation that modifies the size and the charge density distribution.51 This will also result in an increase in the total broadening variance. The second limitation of this method is related to the background noise. Figure 5B displays the Dh,TDA results obtained for the cNP‑30 by using three different sets of PVA capillaries and three-dilution media of different conductivities (BGE, 5-times diluted BGE, and 10-times diluted BGE). The intermediate precision relative standard deviations RSDI (%) are respectively 13.81, 7.26 and 4.13%. In the absence of field amplification during the injection, significantly larger RSDI (13.81%) was obtained. This clearly demonstrates that a low signal-to-noise ratio is also detrimental for the results.

■

Article

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.7b03344. cNP size measurements by DLS in pure water; electrophoretic analysis of the cNPs; effect of the injection time in FASI condition on the electrophoretic profiles of the cNPs; issues related to the TDA analysis of the cNPs; data processing of the Taylorgrams by a halfGaussian fitting; FASI-TDA analysis of the cNPs at different wavelengths (PDF)

■

AUTHOR INFORMATION

Corresponding Author

*Tel.: +33 4 76 63 52 97. Fax: +33 4 76 63 52 98 E-mail: farid. [email protected]. ORCID

Farid Oukacine: 0000-0001-9983-5005 Eric Peyrin: 0000-0001-5558-6369 Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors thank the support of ANR program Labex Arcane (ANR-11-LABX-0003-01). We also acknowledge the NanoBioICMG Platform (FR 2607, Grenoble) for granting access to the Electron Microscopy facility.

■

REFERENCES

(1) Mélinon, P.; Begin-Colin, S.; Duvail, J. L.; Gauffre, F.; Boime, N. H.; Ledoux, G.; Plain, J.; Reiss, P.; Silly, F.; Warot-Fonrose, B. Phys. Rep. 2014, 543, 163−197. (2) Santos, C. S. C.; Gabriel, B.; Blanchy, M.; Menes, O.; Garcia, D.; Blanco, M.; Arconada, N.; Neto, V. Mater. Today Proc. 2015, 2, 456− 465. (3) Salata, O. V. J. Nanobiotechnol. 2004, 2, 3. (4) Bumbudsanpharoke, N.; Choi, J.; Ko, S. J. Nanosci. Nanotechnol. 2015, 15, 6357−6372. (5) Sajid, M.; Ilyas, M.; Basheer, C.; Tariq, M.; Daud, M.; Baig, N.; Shehzad, F. Environ. Sci. Pollut. Res. 2015, 22, 4122−4143. (6) Lichtfouse, E. Sustainable Agriculture Reviews, Vol. 19; Springer International Publishing: Switzerland, 2016. (7) Lin, P. C.; Lin, S.; Wang, P. C.; Sridhar, R. Biotechnol. Adv. 2014, 32, 711−726. (8) Pyrz, W. D.; Buttrey, D. J. Langmuir 2008, 24, 11350−11360. (9) Baalousha, M.; Prasad, A.; Lead, J. R. Environ. Sci. Process Impacts 2014, 16, 1338−1347. (10) Liv, N.; Lazić, I.; Kruit, P.; Hoogenboom, J. P. Ultramicroscopy 2014, 143, 93−99. (11) Jans, H.; Liu, X.; Austin, L.; Maes, G.; Huo, Q. Anal. Chem. 2009, 81, 9425−9432. (12) Schuck, P. Biophys. J. 2000, 78, 1606−1619. (13) Krueger, K. M.; Al-Somali, A. M.; Falkner, J. C.; Colvin, V. L. Anal. Chem. 2005, 77, 3511−3515. (14) Wagner, M.; Holzschuh, S.; Traeger, A.; Fahr, A.; Schubert, U. S. Anal. Chem. 2014, 86, 5201−5210. (15) Striegel, A. M. Anal. Bioanal. Chem. 2012, 402, 77−81. (16) Dorofeev, G. A.; Streletskii, A. N.; Povstugar, I. V.; Protasov, A. V.; Elsukov, E. P. Colloid J. 2012, 74, 675−685. (17) Harada, M.; Saijo, K.; Sakamoto, N.; Einaga, H. Colloids Surf., A 2009, 345, 41−50. (18) Nanomaterial Characterization: An introduction; Tantra, R., Ed.; John Wiley & Sons, Inc.: Hoboken, NJ, 2016.

CONCLUSIONS

In this study, it was demonstrated that the inline coupling of FASI to TDA was a promising characterizing technique for the analysis of low-UV absorbing NPs. This methodology did not require any chromophoric dye to increase the detection sensitivity. An alternative approach to the commonly used Gaussian fitting or integration method was also proposed for the data processing that is based on a half-Gaussian fitting and is particularly adapted for the NPs of narrow size distributions for which the Gaussian shapes of the Taylorgrams are conserved. The R2 values allowing the estimation of the fit goodness can be used to validate this criterion. The obtained Dh values are in very good agreement with those derived from DLS for the stable monodisperse nanoparticles (cNP‑30 and cNP‑50). Intermediate precision relative standard deviation less than 5% were obtained for both NPs. It was demonstrated that the FASI can generate peak dispersion. For such cases, the half-Gaussian fitting method is not adapted for the data processing. Nevertheless this drawback should be overcome by increasing distance between the detection cell and the capillary outlet and by using the integration method. Moreover, a low-to-signalnoise ratio can be detrimental for the Dh,TDA results. G

DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX

Article

Analytical Chemistry

(52) Lim, J. K.; Yeap, S. P.; Che, H. X.; Low, S. C. Nanoscale Res. Lett. 2013, 8, 381. (53) Cho, E. J.; Holback, H.; Liu, K. C.; Abouelmagd, S. A.; Park, J.; Yeo, Y. Mol. Pharmaceutics 2013, 10, 2093−2110. (54) Doncom, K. E. B.; Blackman, L. D.; Wright, D. B.; Gibson, M. I.; O’Reilly, R. K. Chem. Soc. Rev. 2017, 46, 4119−4134. (55) Chamieh, J.; Cottet, H. J. Chromatogr. A 2012, 1241, 123−127. (56) Breadmore, M. C. Bioanalysis 2009, 1, 889−894. (57) Zhang, C. X.; Thormann, W. Anal. Chem. 1996, 68, 2523−2532. (58) Baron, D.; Cacho, C.; Petr, J. J. Chromatogr. A 2017, 1499, 217− 221. (59) Tohala, L.; Oukacine, F.; Ravelet, C.; Peyrin, E. Anal. Chem. 2015, 87, 5491−5495. (60) Harris, K. R.; Woolf, L. A. J. Chem. Eng. Data 2004, 49, 1064− 1069. (61) Sutera, S. P.; Skalak, R. Annu. Rev. Fluid Mech. 1993, 25, 1−19. (62) Poiseuille, J. L. M. Mémoires présentés par divers savants à l’Académie Royale des Sciences de l’Institut de France 1846, IX, 433−544. (63) Hagenbach, E. Ann. Phys. Chem. 1860, 109, 385−426. (64) Feinberg, M.; Lamarque, G. Validation externe des méthodes d’analyse; Les Techniques de l’Ingénieur: France, 2004. Ref : P226 v1. (65) Barbooti, M. M. Environmental applications of instrumental chemical analysis; Apple Academic Press Inc.: Oakville, Canada, 2015.

(19) Hassellöv, M.; Readman, J. W.; Ranville, J. F.; Tiede, K. Ecotoxicology 2008, 17, 344−361. (20) Linsinger, T.; Roebben, G.; Gilliland, D.; Calzolai, L.; Rossi, F.; Gibson, P.; Klein, C. Requirements on measurements for the implementation of the European Commission definition of the term “nanomaterial”; JRC Reference Report EUR 25404 EN; Joint Research Centre: Geel, Belgium, 2012. (21) Hawe, A.; Hulse, W. L.; Jiskoot, W.; Forbes, R. T. Pharm. Res. 2011, 28, 2302−2310. (22) Fissan, H.; Ristig, S.; Kaminski, H.; Asbach, C.; Epple, M. Anal. Methods 2014, 6, 7324−7334. (23) Taylor, G. Proc. R. Soc. London, Ser. A 1953, 219, 186−203. (24) Oukacine, F.; Bernard, S.; Bobe, I.; Cottet, H. J. Controlled Release 2014, 196, 139−145. (25) Latunde-dada, S.; Bott, R.; Hampton, K.; Leszczyszyn, O. I. Anal. Chem. 2015, 87, 8021−8025. (26) Hulse, W.; Forbes, R. Int. J. Pharm. 2011, 416, 394−397. (27) Hulse, W. L.; Forbes, R. T. Int. J. Pharm. 2011, 411, 64−68. (28) Østergaard, J.; Jensen, H. Anal. Chem. 2009, 81, 8644−8648. (29) Sharma, U.; Gleason, N. J.; Carbeck, J. D. Anal. Chem. 2005, 77, 806−813. (30) Cottet, H.; Biron, J. P.; Cipelletti, L.; Matmour, R.; Martin, M. Anal. Chem. 2010, 82, 1793−1802. (31) Holdrich, M.; Liu, S. Y.; Epe, M.; Lammerhofer, M. Talanta 2017, 167, 67−74. (32) Pyell, U.; Jalil, A. H.; Urban, D. A.; Pfeiffer, C.; Pelaz, B.; Parak, W. J. J. Colloid Interface Sci. 2015, 457, 131−140. (33) Balog, S.; Urban, D. A.; Milosevic, A. M.; Crippa, F.; RothenRutishauser, B.; Petri-Fink, A. J. Nanopart. Res. 2017, 19, 287. (34) Høgstedt, U. B.; Schwach, G.; van de Weert, M.; Østergaard, J. Eur. J. Pharm. Sci. 2016, 93, 21−28. (35) Jensen, H.; Østergaard, J. J. Am. Chem. Soc. 2010, 132, 4070− 4071. (36) Latunde-dada, S.; Bott, R.; Hampton, K.; Leszczyszyn, O. I. J. Chromatogr. A 2015, 1408, 255−260. (37) Cottet, H.; Biron, J. P.; Martin, M. Anal. Chem. 2007, 79, 9066− 9073. (38) Cipelletti, L.; Biron, J. P.; Martin, M.; Cottet, H. Anal. Chem. 2015, 87, 8489−8496. (39) Latunde-dada, S.; Bott, R.; Hampton, K.; Patel, J.; Leszczyszyn, O. I. Anal. Methods 2015, 7, 10312−10321. (40) Leclercq, L.; Cottet, H. Anal. Chem. 2012, 84, 1740−1743. (41) Saetear, P.; Chamieh, J.; Kammer, M. N.; Manuel, T. J.; Biron, J. P.; Bornhop, D. J.; Cottet, H. Anal. Chem. 2017, 89, 6710−6718. (42) Hackley, V. A.; Clogston, J. D. Measuring the Size of Nanoparticles in Aqueous Media Using Batch-Mode Dynamic Light Scattering; NIST-NCL Joint Assay Protocol, PCC-1 Version 1.2, U.S. Department of Commerce, 2015; Available at the following: http://dx. doi.org/10.6028/NIST.SP.1200-6. (43) Yamada, H.; Urata, C.; Aoyama, Y.; Osada, S.; Yamauchi, Y.; Kuroda, K. Chem. Mater. 2012, 24, 1462−1471. (44) Huang, X.; Young, N. P.; Townley, H. E. Nanomater. Nanotechnol. 2014, 4, 2. (45) Jäger, E.; Jäger, A.; Etrych, T.; Giacomelli, F. C.; Chytil, P.; Jigounov, A.; Putaux, J.-L.; Ř íhová, B.; Ulbrich, K.; Štěpánek, P. Soft Matter 2012, 8, 9563−9575. (46) Pecora, R. J. Nanopart. Res. 2000, 2, 123−131. (47) Lamberty, A.; Franks, K.; Braun, A.; Kestens, V.; Roebben, G.; Linsinger, T. P. J. J. Nanopart. Res. 2011, 13, 7317−7329. (48) Sprouse, D.; Jiang, Y.; Laaser, J. E.; Lodge, T. P.; Reineke, T. M. Biomacromolecules 2016, 17, 2849−2859. (49) Patterson, J. P.; Robin, M. P.; Chassenieux, C.; Colombani, O.; O’Reilly, R. K. Chem. Soc. Rev. 2014, 43, 2412−2425. (50) Kaasalainen, M.; Aseyev, V.; von Haartman, E.; Karaman, D. Ş.; Mäkilä, E.; Tenhu, H.; Rosenholm, J.; Salonen, J. Nanoscale Res. Lett. 2017, 12, 74. (51) Shaw, R. Dynamic Light Scattering Training; Malvern Instruments: Malvern, U.K., 2014. H

DOI: 10.1021/acs.analchem.7b03344 Anal. Chem. XXXX, XXX, XXX−XXX