Polyacrylamide gel electrophoresis of semiconductor quantum dots and their bioconjugates: materials characterization and physical insights from spectrofluorimetric detection

Hyungki Kim , Tiffany Jeen , Michael V. Tran and W. Russ Algar *
Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, British Columbia V6T 1Z1, Canada. E-mail: algar@chem.ubc.ca

Received 23rd September 2017 , Accepted 18th January 2018

First published on 1st February 2018

Colloidal semiconductor quantum dot (QD) nanocrystals have ideal fluorescence properties for bioanalysis and bioimaging, but these materials must be functionalized with an inorganic shell, organic ligand or polymer coating, and conjugated with biomolecules to be useful in such applications. Several different analytical techniques are used to characterize QDs and their multiple layers of functionalization. Here, we revisit poly(acrylamide) gel electrophoresis (PAGE), which has been scarcely used for the characterization of QDs and their bioconjugates in deference to the routine use of agarose gel electrophoresis. We implemented PAGE in a novel “stubby” capillary format with spectrofluorimetric detection, the combination of which enabled more rapid and more detailed characterization of QDs than was possible with both poly(acrylamide) and agarose slab gels. Correlations between the peak photoluminescence (PL) emission wavelength and electropherogram peaks, especially when combined with Ferguson analysis, provided new and significant insight into the key factors that determine the electrophoretic mobility of QDs, and helped to resolve heterogeneity and sub-populations in ensembles of QDs. The method was useful for characterization of the inorganic core/shell nanocrystals, their organic ligand and polymer coatings, and their final bioconjugates, the latter of which were in the form of peptide and protein conjugates. With further development and optimization, we anticipate that capillary PAGE with spectrofluorimetric detection will become a valuable addition to the toolbox of characterization techniques suitable for QDs, their bioconjugates, and other nanoparticle materials as well.


Colloidal semiconductor quantum dots (QDs) have garnered great interest for applications in bioanalysis and imaging because of their favourable optical properties. These properties include but are not limited to spectrally broad absorption, tunable and spectrally narrow photoluminescence (PL) emission, and orders of magnitude higher brightness and resistance to photobleaching versus conventional fluorescent dyes.1–3 Examples of applications of QDs have included binding assays, Förster resonance energy transfer (FRET)-based assays, optical barcoding, flow cytometry, cellular imaging, intracellular sensing, single-particle tracking, super-resolution imaging, and in vivo imaging, as described in several review articles.4–8 To enable these applications, three layers of chemistry are generally necessary: (i) the core nanocrystal and its protective shell, where the shell is modified with (ii) an organic ligand or polymer coating that imparts colloidal stability and provides chemical handles for conjugation of (iii) biofunctional molecules.9 Achieving the best performance and reproducibility in applications requires optimization of each layer of chemistry, thus making characterization of each layer important.

Many different methods are suitable for characterization of the various layers and properties of QDs and their bioconjugates.10 Optical properties are characterized using methods such as ground-state and transient absorption spectroscopy and steady-state and time-resolved fluorescence spectroscopy.11,12 Physical properties, particularly size and structure, are typically determined by transmission electron microscopy (TEM)13 and X-ray diffraction (XRD),14 although spectroscopic methods are also useful.15–18 Composition is often determined from X-ray photoelectron spectroscopy (XPS),19 energy dispersive X-ray spectroscopy (EDX),20 or inductively coupled plasma mass spectrometry (ICP-MS).21 For the coating layer, infrared absorption spectroscopy can confirm modification of the QD with characteristic functional groups22,23 and, extrapolating from other nanoparticles, thermogravimetric analysis (TGA) and differential scanning calorimetry (DSC) can provide information about the relative amounts of inorganic and organic material.10 Dynamic light scattering (DLS) provides information about hydrodynamic size, inclusive of the nanocrystal, its organic coating, and conjugated biomolecules.24 Analytical ultracentrifugation is similar to DLS in the information it can provide.25 Many of the foregoing techniques are quite sophisticated and specialized, and are frequently accessed only through shared facilities.

Gel electrophoresis is a common characterization technique for QD coatings and bioconjugates that is both simple and readily accessible. This method is widely used to separate and characterize materials based on their size and charge, with the gel acting as a size-selective sieve.26 Agarose and poly(acrylamide) gel (PAG) matrices are most commonly used for the separation and characterization of DNA and proteins. In the case of QDs, agarose gel electrophoresis has been a reliable qualitative and sometimes quantitative method for characterizing the modification of QDs with ligand or polymer coatings and conjugated biomolecules.24,27,28 The experiments are quick, comparatively low-cost, and simpler to execute and analyze than the other characterization methods noted above. Although PAGs offer more uniform pore sizes and higher resolution, the characterization of QDs has almost exclusively relied on agarose gels, presumably because of their ease of preparation and larger pore size. There are very few reports on the use of PAGs with QDs, and these reports have featured only qualitative analysis and generally shown very low mobility, significant streaking, and otherwise poor visualization.29–32

Here, we revisit the use of poly(acrylamide) gel electrophoresis (PAGE) as a method for the physical characterization of QDs and their bioconjugates. PAGE is implemented in a “stubby” capillary format with spectrofluorimetric detection to gain insight into the factors that affect migration of the QDs. The “stubby” format, defined by the short length and relatively large diameter of the capillary used, represents a compromise between classic slab PAGE and conventional capillary PAGE. Some of the advantages of this format are a footprint small enough to be mounted on the stage of a microscope, low sample consumption, higher field strengths at lower voltages, and better resolution and shorter run times versus agarose and poly(acrylamide) slab gel electrophoresis. We evaluated QD materials that varied in the properties of the inorganic semiconductor nanocrystal, the nature of the organic ligand or polymer coating, and the degree of conjugation with peptide and protein. Trends in PL emission wavelength were compared with features in electropherograms to gain insight into the key factors that determined electrophoretic mobility, and to help resolve heterogeneity and sub-populations in ensembles of QDs. With further optimization and development, PAGE with spectrofluorimetric detection has the potential to be an excellent tool for the characterization of QDs and their bioconjugates.

Experimental section

Additional details can be found in the ESI.

QD materials and coatings

CdSe/CdS/ZnS QDs were synthesized using standard hot solvent methods33,34 and are denoted as X-QDλ, where λ is their peak emission wavelength (in nm) and X is their surface chemistry. The QDs were dispersed in water through the X surface chemistry, either via ligand exchange or overcoating with a polymer. Ligand exchanges were done with dihydrolipoic acid (DHLA), DHLA-sulfobetaine (DHLA-SB), and glutathione (GSH). Polymer coatings comprised a poly(isobutylene-alt-maleic anhydride) backbone conjugated with either hexadecylamine (HDA-PMA) or 1-(3-aminopropyl)imidazole (API-PMA). Full details can be found in the ESI.

A 3.1 kDa peptide (Biosynthesis Inc., Lewisville, TX, USA; GGNGSGQNGAAYALVPRGSG-P5GH6) and recombinant Protein A (SPA; ab52953, Abcam Inc., Toronto, ON, Canada) were assembled on QDs through hexahistidine tags, which is a well-established method for the conjugation of biomolecules to QDs.35,36 Peptides were mixed with QDs at room temperature for ∼2 h prior to measurements, and proteins were mixed with QDs for ≥12 h prior to measurements.

Preparation of poly(acrylamide)-filled capillaries

The inner walls of capillaries were first modified with vinyl groups. Borosilicate glass capillaries (75 mm length, 0.4 mm inner diameter; Drummond Scientific Co., Broomall, PA, USA) were sonicated in ethanol for 2 h, drained, dried in air, plasma cleaned for 45 s (Harrick Plasma PDC-32G; Ithaca, NY, USA), incubated for >4 h in 2% v/v solution of 3-(trimethoxysilyl)propyl methacrylate (TMSM) or allyltrichlorosilane (Sigma-Aldrich, Oakville, ON, Canada) with 1% v/v glacial acetic acid, drained, and then dried in air in an oven at ∼70 °C for >1 h. Both silane modifications were equally effective for the preparation of PAG-filled capillaries; however, TMSM was used in most experiments.

PAGs were prepared by diluting a 40% w/w solution of 37.5[thin space (1/6-em)]:[thin space (1/6-em)]1 acrylamide[thin space (1/6-em)]:[thin space (1/6-em)]bis-acrylamide solution in borate buffer (100 mM, pH 9.2) to a final volume of 1.49 mL. Tetramethyl-ethylenediamine (TEMED; 1.5 μL) and 25% w/v ammonium persulfate (aq) (APS; 4.5 μL) were then added, and the final solution was vigorously mixed in a microcentrifuge tube. Different percentages of PAGs were prepared by scaling the amount of acrylamide[thin space (1/6-em)]:[thin space (1/6-em)]bis-acrylamide solution accordingly. A vinyl group-modified glass capillary was inserted through a small hole in the lid of the microcentrifuge tube, filled by capillary action, and left at room temperature for 2 h. Gel-filled capillaries were soaked in buffer for >20 min prior to experiments to help quench residual radicals. Unless stated otherwise, 3.0% w/w gel was used in experiments. Fig. 1 summarizes the chemistry for glass modification and PAG gel formation.

image file: c7an01581j-f1.tif
Fig. 1 Treatment of a glass capillary with TMSM to yield vinyl groups for covalently immobilizing PAGs. The structure of TMSM has been truncated for clarity. The R group in the figure is one of several possible outcomes of radical polymerization that will be found in a PAG.


Fig. 2 shows the design of the apparatus for spectrofluorimetric detection of QDs migrating through PAG-filled capillaries. The capillary was supported on a glass microscope slide that had a hole drilled out to allow the microscope objective to focus on the capillary. Buffer reservoir zones were created by application of a hydrophobic coating (Sigmacote; Sigma-Aldrich) to all other areas of the slide. The capillary spanned the distance between the two reservoirs, which held ∼0.65 mL of borate buffer (100 mM, pH 9.2) as the running buffer. Platinum wire electrodes attached to a high-voltage (HV) sequencer (ER230 HV sequencer, eDAQ, Colorado Springs, CO, USA) were immersed in both buffer reservoirs. The microscope slide was held in custom-built clear Plexiglas (poly(methyl methacrylate)) housing with external HV connectors and internal post terminals, and a lid with a magnetic safety interlock.
image file: c7an01581j-f2.tif
Fig. 2 (A) Schematic of the apparatus. The Plexiglas housing (1) has external HV connectors for the anode (2) and cathode (3), internal posts (4) for attaching Pt wires (5), and holds a 25 mm × 75 mm glass slide (6) that has a hole drilled out to permit approach of the objective lens (7) of a microscope. The slide is further patterned with a hydrophobic coating to create two buffer reservoirs (8) that the glass capillary (9) spans. Bands of fluorescent materials (10) are detected as they migrate through the focus of the objective lens. The direction of migration of QDs and fluorescein is shown by the arrow. (B) Photograph of the apparatus mounted on an inverted microscope stage. (C) Simplified schematic of the optical path for real-time spectrofluorimetric detection, including excitation light (11), objective lens (7), dichroic mirror (12), mirrors (13), longpass filter (14), lens (15) and optical fibre connection (16) to a portable CCD spectrometer (17).

For spectrofluorimetric detection, the apparatus was mounted on the translation stage of an inverted microscope (IX83, Olympus, Richmond Hill, ON, Canada) equipped with a metal–halide light source (Excelitas Technologies, Mississauga, ON, Canada), dichroic mirror (cut-off wavelength 470 nm; Chroma Technology Corp, Bellow Falls, VT, USA), longpass filter (cut-off wavelength 530 nm; Thorlabs Inc., Newton, NJ, USA), and connection to a fibre-optic spectrometer (GreenWave 16 VIS-50, StellarNet, Tampa, FL, USA). The fiber-optic connection was a round-to-linear bundle (7 × 200 μM diameter core fibres; Thorlabs). PL emission spectra as a function of migration time were recorded using custom software written in LabVIEW (National Instruments, Austin, TX, USA).

Capillary gel electrophoresis

Fluorescein (final concentration between 1–25 μM) was added to each QD sample to serve as an internal standard for comparison of migration times between electropherograms. Samples were electrokinetically injected into the capillary using an HV sequencer at 300 V for 5 s. The sample solution was then removed and replaced with running buffer. Capillary PAGE runs were then completed at 300 V for a field strength of ∼55 V cm−1. The QD concentrations in sample solutions for PAGE experiments were 0.5 μM (QD520, QD570), 0.2 μM (QD600), and 0.1 μM (QD650). The distance from the start of the capillary to the detection point was ∼4.2 cm.

Ferguson analysis

Ferguson plots were analyzed as per eqn (1), where the mobility, M, was calculated as the quotient of the migration velocity (cm s−1) and field strength (V cm−1), M0 was the extrapolated free mobility of the particle in the absence of the gel, KR was the retardation factor, and T was the gel concentration.24,37,38 The gel concentration was varied between 3.0% and 6.0% while keeping the acrylamide[thin space (1/6-em)]:[thin space (1/6-em)]bisacrylamide ratio at 37.5[thin space (1/6-em)]:[thin space (1/6-em)]1.
log[thin space (1/6-em)]M = log[thin space (1/6-em)]M0KRT(1)


Effect of PAG density

Our first set of experiments was to test the effect of different PAG densities on the migration of QDs. For this purpose, GSH-QD650 were used as a model material with PAG concentrations between 3.0% and 6.0%. At less than 3.0% concentration, poly(acrylamide) did not gel reproducibly. Fig. 3A shows that the migration time increased for the QDs as the gel concentration increased. This trend was expected with decreasing pore size as the gel density increased. The electropherogram peak full-width-at-half-maximum (FWHM) increased with increasing gel density, and peak asymmetry also increased in the form of greater tailing up until 6.0%, when fronting became significant. Spectrofluorimetric detection enabled an analysis of the QD PL emission at different points in each of the electropherograms, as shown in Fig. 3B for 3.0% PAG. The wavelength of maximum PL emission was shorter at the leading edge of the electropherogram peak and gradually red shifted toward the trailing edge. The spectrum at the maximum of the electropherogram peak was a good match to the PL spectrum measured for the ensemble of QDs in bulk solution, albeit slightly more narrow in spectral FWHM. Analogous results were observed at other gel densities, as shown in Fig. 3C, which plots the peak PL emission wavelength as a function of time and gel density. In contrast, the peak PL emission wavelength for the fluorescein internal standard was approximately constant. The signal-to-noise ratio for spectrofluorimetric detection gradually decreased with increasing gel density and became impractical at gel densities greater than 6.0%. PAGs with 3.0% density were used for further experiments because they had the best signal-to-noise ratios, the fastest runs, and most narrow electropherogram peaks.
image file: c7an01581j-f3.tif
Fig. 3 (A) PL electropherograms of GSH-QD650 at different PAG densities. All other conditions were kept identical. (B) PL emission spectra at selected points (circled) in the electropherogram peak for QD650 with a 3.0% PAG. The shaded spectrum is the ensemble PL spectrum measured in bulk solution. Spectra for other densities can be found in the ESI. (C) Changes in the peak PL emission wavelength as a function of migration time. The legend in panel A also applies to panel C. Note that the time axis has the same scale between panels A and C. The horizontal dashed line is the wavelength of peak PL emission for the GSH-QD650 in bulk solution. The vertical dashed lines indicate the electropherogram peak positions.

Series of quantum dots

In the next set of experiments, batches of QDs with different PL emission colours were compared on 3.0% PAG capillaries and an agarose slab gel. These batches were QD520, QD570, QD600, and QD650. Some relevant properties of each batch of QDs are summarized in Table 1.16Fig. 4A and B show a PL image and PL intensity profile of an agarose slab gel with GSH-coated preparations of each batch of QDs. The QD520 had the greatest electrophoretic mobility whereas the QD570, QD600, and QD650 had similar mobilities. Fig. 4C shows PL electropherograms for the same preparations of QDs with PAGE. The QD520 once again had the highest mobility and, with PAGE, differences in the migration time and peak shape between the other batches of QD were more clearly resolved. Fig. 4D plots the peak PL emission wavelength versus migration time for each of the electropherogram peaks in Fig. 4C. As with the QD650, a gradual bathochromic shift in the peak PL emission was observed with increasing migration time for QD600, QD570, and QD520. The magnitude of the shift for the QD520 was artificially diminished by spectral overlap with the fluorescein standard early in its peak and by partial cropping of its full PL emission spectrum by the longpass filter in the apparatus. The latter also affected the QD570, albeit to a lesser extent.
image file: c7an01581j-f4.tif
Fig. 4 (A) PL image and (B) corresponding PL intensity profile of an agarose gel with four different batches of GSH-QDλ. The faint band for the QD570 in panel A is outlined for clarity. (C) PL electropherograms of the same GSH-QDs with a 3.0% PAG. (D) Changes in the peak PL emission wavelength as a function of migration time for the four batches of GSH-QDλ analyzed by PAGE. The legend in panel C also applies to panels B and D. The horizontal dashed lines are the wavelengths of peak PL emission for the QDλ in bulk solution. The vertical dashed lines indicate the electropherogram peak positions. No horizontal line is shown for the QD520 because the longpass filter used for measurements distorted the spectrum (see ESI for details).
Table 1 Selected properties of different batches of GSH-QDλ
Property QD520 QD570 QD600 QD650
All values are ± 1 standard deviation for a minimum of three replicates.a Inclusive of the core and shell. Estimated by measurement of 50 individual QDs.b From the ground state absorption spectrum measured in bulk solution.c Estimated from the method of Yu and Peng (ref. 16).d From the PL emission spectrum measured in bulk solution.e The migration time is adjusted so that the fluorescein electropherogram peak is at 0 min.
TEM size (nm)a 5.4 ± 1.4 6.0 ± 0.8 9.8 ± 1.3 8.7 ± 0.9
1st Exciton peak (nm)b 496 552 584 632
Approximate core diameter (nm)c 2.3 3.1 4.0 6.4
PL emission maximum (nm)d 521 570 602 645
PL emission FWHM (nm)d 36 30 32 28
Adjusted migration time in 3.0% PAG (min)e 0.79 ± 0.02 1.82 ± 0.06 2.08 ± 0.05 1.83 ± 0.04
Electropherogram peak FHWM (min) 1.37 ± 0.02 1.9 ± 0.1 1.4 ± 0.2 0.57 ± 0.02
Electropherogram peak asymmetry 5.1 ± 0.2 2.3 ± 0.3 4.3 ± 0.9 2.0 ± 0.1

Ligand and polymer coatings

All experiments to this point assessed the electrophoretic mobility of GSH-QDs. Our next step was to evaluate different coatings on a QD by capillary PAGE. To this end, QD650 were also coated with anionic DHLA ligands, zwitterionic DHLA-SB ligands, anionic HDA-PMA amphiphilic polymer (two batches), and anionic API-PMA coordinating polymer. Fig. 5A shows the structures and putative morphology of the coated QDs, and Fig. 5B shows representative electropherograms for these X-QD650 when run on a 3.0% PAG. The migration time followed the trend GSH-QD ≈ DHLA-QD < API-PMA-QD < HDA-PMA-QD (batch 1) < DHLA-SB-QD, where the HDA-PMA-QD (batch 2) did not have an unambiguous migration time. The first batch of HDA-PMA-QD had a well-defined peak but also had a pronounced tail, whereas the second batch appeared to have two leading peaks (local maxima) that were overwhelmed by a larger tail. For both preparations, the most mobile of the HDA-PMA-QD migrated at rates that were slightly slower than the API-PMA-QD. The FWHM for the first batch of HDA-PMA-QD was similar to that for the API-PMA-QD, but could not be defined for the second batch. Variations in the overall effectiveness and aqueous dispersion yield of amphiphilic polymer coating methods are well known,39 so the differences between the two batches of HDA-PMA-QD were not unexpected, albeit not intentional.
image file: c7an01581j-f5.tif
Fig. 5 (A) Structures (i) and putative coating morphologies (ii) for GSH, DHLA, DHLA-SB, HDA-PMA, and API-PMA on QDs. In (ii), the ligands are drawn as line structures with coloured circles for heteroatoms, whereas the polymer coatings are drawn as cartoons with no structure for the side chains and backbone. The red circles on the HDA-PMA and API-PMA cartoons are carboxyl groups, and the blue circles on API-PMA are imidazole groups. (B) Representative PL electropherograms of X-QD650, where X is a ligand or polymer. (C) Changes in the peak PL emission wavelength as a function of migration time for the electropherogram peaks in panel B. The horizontal dashed lines are the wavelengths of peak PL emission for the X-QDs in bulk solution. The vertical dashed lines indicate the electropherogram peak positions. The legend in panel B also applies to panel C; however, the time axes scale differently between the panels. (D) Ferguson plots for X-QD650, where X = GSH, DHLA-SB, API-PMA, and HDA-PMA (batch 2), and the PAG concentration varies between 3.0 and 6.0%. A plot for fluorescein is also shown for reference.

Fig. 5C tracks the peak PL emission wavelength over time for the electropherogram peaks for each X-QD650. As before, the GSH-QD exhibited a PL emission peak shift of ∼8 nm over the width of its electropherogram peak. The DHLA-QD were very similar with a PL emission peak shift of ∼11 nm. Curiously, the DHLA-SB-QD exhibited a much smaller PL emission peak shift of only ∼4 nm and started at a longer wavelength at the leading edge of the electropherogram peak. Interesting results were also obtained with the API-PMA-QD and the HDA-PMA-QD. The API-PMA-QD exhibited a gradual PL emission peak shift of ∼3.5 nm across the electropherogram peak, which then became constant over the tailing shoulder of the electropherogram peak. The first and second batches of HDA-PMA-QD exhibited an initial shift of ∼8 nm in the peak PL emission, and this increase aligned with the first apparent peak in the electropherogram (indicated by the single asterisk). For the second batch, a blip in the wavelength trend was then observed and appeared to correspond to the second peak in the electropherogram (indicated by the double asterisk). As with the API-PMA-QD, the peak PL emission wavelength became constant for both batches of HDA-PMA-QD in the tailing regions.

Fig. 5D is a Ferguson plot analysis of four of the six different X-QD650 and a fluorescein control. DHLA-QDs were not analyzed because their mobilities on agarose (≤1.5% w/w) and PAG (3.0% w/w) were indistinguishable from GSH-QDs. The second batch of HDA-PMA-QD was analyzed instead of the first batch in order to see the impact of gel density on the apparent polydispersity. The first peak in its electropherogram was used as the migration time. Table 2 summarizes the results from the Ferguson analysis. As expected, the change in PAG density affected the fluorescein far less than the QDs. The slopes of the Ferguson plots and thus the retardation factors for the GSH-QD and DHLA-SB-QD were indistinguishable within experimental error; however, the DHLA-SB-QD had a smaller free mobility and zeta potential. In turn, the API-PMA-QD had a free mobility and zeta potential indistinguishable from the GSH-QD, but had a larger retardation factor. The HDA-PMA-QD (batch 2) had the highest free mobility and zeta potential, as well as the highest retardation factor. As shown in the ESI, increases in PAG density gradually increased the relative magnitude of the first and second peaks versus the tailing. Estimated hydrodynamic radii for the various X-QD650 were determined from fitting the Ferguson plot data (see ESI for details).

Table 2 3.0% PAG and Ferguson analysis results of selected X-QD650a
X-QD650 3.0% adj. migration timec (min) 3.0% FWHM (min) Effective radius (nm) Zeta potential (mV)
a All values are ± 1 standard deviation for a minimum of three replicates. nd = not determined; na = not applicable. b Values for the first, second maxima in the electropherogram. c The migration time is adjusted so that the fluorescein electropherogram peak is at 0 min.
X = GSH 1.83 ± 0.04 0.57 ± 0.02 3.6 ± 0.6 −76
X = DHLA-SB 7.21 ± 0.08 1.4 ± 0.2 3.3 ± 0.6 −49
X = API-PMA 4.4 ± 0.1 1.9 ± 0.3 5.1 ± 0.5 −70
X = HDA-PMA 1 4.94 ± 0.05 1.83 ± 0.03 nd nd
X = HDA-PMA 2b 5.7 ± 0.5, 7.4 ± 0.5 na 16 ± 1, 15 ± 1 −233, −171


Having analyzed various QDs and their coatings, the next analyses were QDs conjugated with peptide and protein molecules. The peptide sequence was 32 amino acid residues (∼3.1 kDa) and had a hexahistidine tag at its C-terminus. The protein was a recombinant Staphylococcal protein A (SPA; ∼39 kDa) with a hexahistidine tag at its N-terminus. In both cases, the role of the hexahistidine tag was to bind to the ZnS shell of the QDs, as has been thoroughly characterized and utilized in the literature.35,36Fig. 6A shows a PL image of an agarose slab gel with GSH-QD600-[peptide]N and GSH-QD600-[SPA]N conjugates, where N is the average number of peptide or protein equivalents per QD across the ensemble. As expected from previous studies,24,27,28,40 the QD600-[peptide]N conjugates remained as a single band that gradually decreased in mobility with increasing N, and the QD600-[SPA]N conjugates split into multiple bands at low N until converging into a single band of low mobility at large N. PL profiles for the QD600-[peptide]N and QD600-[SPA]N conjugates on the agarose gel are also shown in Fig. 6A.
image file: c7an01581j-f6.tif
Fig. 6 (A) PL image and profiles for an agarose slab gel with GSH-QD600-[peptide]N and GSH-QD600-[SPA]N conjugates, where N is the average number of peptide or protein equivalents per QD across the ensemble. (B) PL electropherograms and corresponding changes in the peak PL emission wavelength over time for the same GSH-QD600-[peptide]N on a 3.0% PAG. The horizontal dashed line is the wavelength of the peak PL emission for the QD600 in bulk solution. The vertical dashed lines indicate the electropherogram peak positions. (C) Data analogous to panel B for GSH-QD600-[SPA]N with 3.0% PAGE.

Fig. 6B shows PL electropherograms and the corresponding trends in the peak PL emission wavelength for the QD600-[peptide]N conjugates analyzed by capillary PAGE. The same trend in migration time was observed between the capillary PAG and the agarose slab gel, although the N = 16 and N = 32 conjugates were better resolved with the PAG. Each conjugate exhibited a steady increase in its peak PL emission wavelength across the electropherogram peak. Fig. 6C shows analogous PAGE data for the QD600-[SPA]N conjugates. Similar to the multiple bands on an agarose slab gel, multiple electropherogram peaks were observed for QD600-[SPA]N conjugates on a PAG; however, the pattern of the peaks was different. As N increased beyond zero, the original peak decreased and a second and lower mobility peak grew; in turn, the second peak decreased and a third and even lower mobility peak grew. This increase in the number of peaks and shift in intensity to lower mobility peaks continued up to four or five peaks for N = 8, where the putative fifth peak was a shoulder on the fourth peak and not fully resolved. (The N = 16 sample did not produce adequate signal-to-noise in PAGE for analysis.) Resolution of the peaks for N = 0 through N = 4 was greater with PAG than with agarose. Unlike the peptide data, the trend in the peak PL emission wavelength for the protein conjugates was non-monotonic. Undulations in wavelength were observed in parallel with each peak in the electropherogram, diminishing in magnitude with each successive peak.


The role of nanocrystal size polydispersity

The electrophoretic mobility of a nanoparticle in a gel is a function of its size and charge. Given that a QD can be approximated as a sphere with a discrete number of charges on its surface, there is an inherent competition between tendencies for (i) decreased mobility with an increase in the size of the QD nanocrystal, and (ii) an increased mobility with the concurrent increase in surface area and thus capacity for bound charge from a ligand or polymer coating. Parallel measurement of electropherograms and PL emission spectra provided insight into this competition because the peak PL emission wavelength of a QD is dependent on the size of its core nanocrystal.

The electrophoretic mobility of a QD in a sieving medium such as PAG or agarose gel is approximated by the aforementioned eqn (1), which can be rewritten as eqn (2) when the retardation factor is given by eqn (3).24,37 The terms a and b in eqn (3) are empirical constants and R is the effective radius of the QD.

log[thin space (1/6-em)]M = log[thin space (1/6-em)]M0 − (a2R2 + 2abR + b2)T(2)
image file: c7an01581j-t1.tif(3)

In the simplest model,41 the free mobility of a spherical insulating particle is given by eqn (4), the Hückel equation, which is the approximation that we apply to QDs. The terms in this equation are the viscosity of the medium, η, the relative permittivity, εr, the vacuum permittivity, ε0, and the particle zeta potential, ζ.

image file: c7an01581j-t2.tif(4)

In turn, the simplest model for the zeta potential is eqn (5),41 where qs is the surface charge. We then make the assumption in eqn (6) that the surface charge is directly proportional to the surface area of the QD for a particular type of ligand coating, where AL is the average footprint of a bound ligand with a charge of −1, and e is the elementary charge.

image file: c7an01581j-t3.tif(5)
image file: c7an01581j-t4.tif(6)

Through substitutions, eqn (3)–(6) lead to eqn (7), which shows that the electrophoretic mobility of a QD should decrease with increasing R because the retardation term increases faster than the free mobility term.

image file: c7an01581j-t5.tif(7)

With the assumption that shell thickness is uniform across the ensemble of QD cores, the qualitative expectation of eqn (7) is confirmed by the observed trends in the peak PL emission wavelength as a function of migration time for ligand-coated QDs. Size polydispersity is therefore a main contribution to the electropherogram peak width for these QD materials.

Assessing the heterogeneity of X-QDs

The ability to track the shift in the peak PL emission wavelength of QDs across their electropherogram peak is a basis for resolving heterogeneity and sub-populations in an ensemble of X-QDs. For example, the uniformity of shell thicknesses across the ensemble of core nanocrystals can be assessed because the core size determines the emission wavelength but both the core and the shell determine the overall size of the QD. Variation in shell thickness, if non-correlated with core size, will reduce the magnitude of the wavelength shift across the electropherogram peak that would otherwise be caused by core size polydispersity. The assumption in this analysis is that the organic coating is uniform. It follows that the converse analysis is also possible: if shell thickness is uniform, then the homogeneity of the organic coating can be assessed. Overall, the large shifts in peak PL emission wavelength for our X-QD coated with X = GSH and DHLA indicate good homogeneity in both shell thickness and coating.

Despite the above, there were still indications of heterogeneity in the ensemble of QDs, albeit seemingly unrelated to the polydispersity of the core nanocrystal. The trends in peak PL emission wavelength with migration time in Fig. 3–6 showed curvature and a plateau (or sometimes a reversion) over the electropherogram peak tails. As explained in the ESI, an approximately linear trend was expected for only polydispersity in the core size. The discrepancy suggested a source of heterogeneity other than core size. One possibility was variation in shell thickness across the ensemble of core QDs; a second possibility was sub-populations of QDs with less than a saturating number of ligands (i.e. charges), where these sub-populations would have had the same mobility as larger QDs with a saturating number of ligands. The observed PL emission maximum was an average of all QDs with the same mobility, inclusive of combinations of core size, shell thickness, and ligand numbers that led to the same net mobility. We therefore suggest that heterogeneity in the latter two properties led to the non-linear increase and eventual plateau or reversion in peak PL emission wavelength with migration time.

Although not entirely homogeneous, the large shifts in peak PL emission wavelength across the electropherogram peak suggested that the GSH- and DHLA-QD650 ensembles were nonetheless good reference materials against which to compare the other X-QD650. For example, the API-PMA-QD650 exhibited a smaller shift in peak PL emission wavelength than the GSH- and DHLA-QD650, and the FHWM of its electropherogram peak was larger. These two results suggested greater heterogeneity and a source other than the inorganic component of the QD. By deduction, this source must have been the API-PMA coating, which was polydisperse in its molecular weight and able bind to QDs at multiple points in its structure. The data may thus indicate multiple binding modes. The Ferguson analysis estimated the effective hydrodynamic radius of the API-PMA-QD650 to be 4.8 nm, a value that was 1.5 nm larger than that for GSH-QD650, consistent with binding modes where the API-PMA polymer was not wrapped tightly around the QD. An effective radius similar to GSH-QD650 would have been expected if the polymer wrapped around the QD perfectly tight. Interestingly, a small shoulder was observed on the late edge of the main peak for API-PMA-QD650, which suggested the potential for a sub-population of QDs. When the PAG density was increased from 3.0% to 3.5%, this shoulder became a defined peak. As the PAG density increased further, this trailing peak separated from the main peak further, broadened and diminished in intensity until vanishing at 6.0% PAG (see Fig. S5). At low PAG densities, the peak PL emission wavelength gradually reverted to the ensemble average across the shoulder/trailing peak, but was approximately constant at higher PAG densities. These results suggested a species that was larger than the individual API-PMA-QD650, sufficiently well-defined to give a peak, and had a mobility that was not a direct function of core nanocrystal size. Dimers of API-PMA-QD650 fit this description. The combination of two QD650 with different core sizes would have a net PL emission wavelength that would tend back toward the ensemble average or shorter wavelengths if dimers of smaller QDs led small-large and large-large dimer combinations.

Similar to the API-PMA-QD650, the DHLA-SB-QD650 had a smaller shift in the peak PL emission wavelength and larger FHWM versus GSH-QD650 and DHLA-QD650. Unlike API-PMA, DHLA-SB is not a polymer but a small molecule ligand. The greater heterogeneity associated with the DHLA-SB-QD650 versus DHLA-QD650 must have therefore arisen from the differences in the number, arrangement, and (potentially) purity of the ligands. Differences between the ligand exchange procedures for DHLA-SB and DHLA included the method of reduction of the ligand, solvent composition, reaction time and temperature—all of which may have contributed to a difference in the number and arrangement of ligands at the QD interface. The Ferguson analysis estimated a very similar effective hydrodynamic radius versus the GSH-QD650 but a lower free mobility and zeta potential for the DHLA-SB-QD650. This size similarity was expected given the sizes of the ligands and their well-defined binding mode, whereas the zeta potential difference presumably arose from the zwitterionic character of the DHLA-SB ligand. A small amount of tailing of the DHLA-SB-QD650 electropherogram peak was observed and did not exhibit any trend in the peak emission wavelength nor evolve into a better defined shoulder or peak with increases in PAG density (see Fig. S4). This result suggested a small but heterogeneous fraction of the ensemble that may have had less ligand coverage resulting in fewer DHLA-SB ligands per QD than the majority.

Turning to HDA-PMA-QD650, the first batch of these QDs had an obvious electropherogram peak, albeit with substantial tailing. The trend in the peak PL emission wavelength for the first batch exhibited a monotonic increase over the peak with no trend over the tail. In contrast, the electropherogram for the second batch of HDA-PMA-QD650 was drastically different than those for all other X-QD650 materials in that it did not have a well-defined peak. Overall, the “peak” was equivalent to a smear on a slab gel, although the leading edge appeared to comprise two discrete sub-populations of QDs. These sub-populations were indicated by both local maxima in the electropherogram and the trend in the peak PL emission wavelength, the latter of which undulated with the two maxima, albeit with lower amplitude for the second maximum. The peak PL emission wavelength plateaued after the second maximum, suggesting that no further discrete populations were resolved. Notably, the position of the first maximum in the electropherogram for the second batch of HDA-PMA-QD650 aligned with the well-defined peak for the first batch of these QDs. Ferguson analysis of the second batch of HDA-PMA-QD650 estimated roughly equal hydrodynamic radii but slightly different zeta potentials between the first and second local maxima in the electropherogram. To account for these results, the total charge on the two sub-populations of QD must have been different. One possibility is that both sub-populations were individual QDs coated in polymer, where the second population had fewer polymer chains per QD without a significant decrease in size. The tail of electropherogram was too heterogeneous to interpret, particularly for the second batch. Sources of heterogeneity may have included large variation in the number of polymer chains per QD and perhaps dimers of QDs. Larger sizes for the material(s) in the peak tail was confirmed by the progressively diminished contribution to the electropherogram profile as PAG density increased.

The migration of QD-bioconjugates

As with the X-QDs, correlations between electropherogram and peak PL emission wavelength data enabled deeper insight into the migration and characterization of GSH-QD600-[peptide]N and GSH-QD600-[SPA]N conjugates. It was possible to assess effects from and differences between the peptides and SPA because the underlying GSH-QD600 exhibited a large shift in peak PL emission wavelength across the electropherogram.

The electropherogram peak for GSH-QD600-[peptide]N conjugates increased in migration time, FWHM, and symmetry as N increased, and there was a large shift in peak PL emission wavelength for each N. Although these shifts would suggest good homogeneity, single particle FRET and gel electrophoresis data have shown that the hexahistidine-tagged peptides and proteins assemble to QDs with stoichiometry that follows a Poisson distribution.42 The QD-[peptide]N conjugates were therefore not homogeneous. Instead, the PAGE results can be rationalized in terms of the flexibility of the peptides and their ability to adopt many conformations as the QD conjugates migrate through the pores of the gel. This range of conformations translated into a range of electrophoretic mobilities that convolved with the different mobilities of conjugates with different values of N. The inflexible size of the QD was thus able to retain a non-trivial contribution to the overall electrophoretic mobility while the increasing contribution from the peptide number and conformational freedom accounted for the gradual increase in migration time, the gradual loss of peak asymmetry, the gradual increase in FWHM, and the gradual decrease in the magnitude of the wavelength shift (∼18 nm for N = 0 with a somewhat incremental decrease to ∼12 nm for N = 32).

Compared to the peptide, SPA was less flexible and less dynamic in its conformation as a folded protein, and it was more than an order of magnitude larger by molecular weight. The SPA therefore had a greater and less variable effect on the mobility of the QDs. Multiple peaks that corresponded to each incremental increase in the number of SPA molecules per QD were observed in the electropherogram. The pattern of peaks should have followed a Poisson distribution, but appeared to have been skewed to lower values than expected because of a size exclusion limit (vide infra). That each peak was a discrete population was clearly indicated by the undulations in the peak PL emission wavelength. These undulations decreased in magnitude as the number of SPA per QD increased, reflecting the progressively larger contribution of the protein to the overall electrophoretic mobility of the conjugate. Although the protein was folded and had a well-defined attachment point to the QD, there would have nonetheless been heterogeneity in where (e.g. different nanocrystal facets) and how (i.e. some limited degrees of freedom) each successive protein molecule attached. At larger N, this protein heterogeneity dominated over the contributions of the QD600 to the net electrophoretic mobility.

Interpretation of spectral PAGE data

Given the data discussed above, we propose the following guidelines for interpretation of spectral PAG electrophoresis data with QDs and their bioconjugates: (i) the larger and more linear the increase in peak PL emission wavelength across an electropherogram peak, the more homogeneous the population of QDs in terms of shell thickness and ligand or polymer coating; (ii) undulations in the peak PL emission wavelength are good indicators of discrete and well-defined sub-populations of QDs when correlated with peak or shoulder features in the electropherogram; (iii) irregular features in the electropherogram without any trend in peak PL emission wavelength suggest substantial heterogeneity; and (iv) the effects of the number, size, shape, and flexibility of conjugated biomolecules must be considered and may introduce heterogeneity that masks homogeneity in the shell thickness and coating of the underlying QD.

Poly(acrylamide) versus agarose

An important result with the PAGE was that the trends in the electrophoretic mobilities of the QD materials mirrored those with an agarose gel, both in terms of the migration time and the peak/bandwidths. “Stubby” capillary PAGE was thus able to access the same information as a slab agarose gel while offering several other advantages. In particular, the “stubby” capillary format facilitated the setup for spectrofluorimetric detection (the benefits of which were discussed above) and the format also had lower sample consumption, higher electric field strength, faster run times, and greater resolution than slab agarose gels. It was not possible to run the experiments with agarose in the capillary because it extruded under the applied field, as reported elsewhere.43 Although PAG did not extrude, we nonetheless found it beneficial to modify the capillary wall with vinyl groups that covalently linked the PAG to the capillary during polymerization. This modification improved reproducibility and made it possible to use a typical PAG capillary for up to ∼10 runs before a new one was needed.

The PAGE was not without some disadvantages. Although the run times were faster, the total preparation time for a PAG-filled capillary was longer than the preparation time for an agarose slab gel. Fortunately, less reagent was consumed and it was possible to prepare the capillaries in batches and store them in buffer. Net neutral QDs were also unsuitable for analysis as there was no means of detecting the absence of mobility, unlike a slab gel, which can be imaged. Another limitation was an apparent size exclusion limit, as suggested by the results of two experiments. First, for the same concentration of QDs, the PL signal-to-background ratio decreased significantly as the PAG density increased, suggesting that fewer QDs were making it to the detection point. Second, the pattern of peaks for the GSH-QD600-[SPA]N conjugates differed significantly between PAG and agarose. The relative intensities of the PAG electropherogram peaks for higher values of N were much lower than expected based on the band intensities observed with the agarose gel. Moreover, the absolute intensities for all peaks also decreased as N increased. These discrepancies suggested the exclusion of conjugates with more than five protein molecules per QD, although the exclusion limit did not appear sharp. A fraction of larger conjugates likely migrated to the detection point through pathways with larger pore sizes, whereas other conjugates were trapped by smaller pores. Transient trapping of QDs in heterogeneous PAGs has been previously observed by single particle tracking44 and supports this hypothesis of a soft or gradual exclusion limit.

Further to the exclusion limit, we observed a curious result when analyzing QD-[peptide(dye)]N conjugates, where the dye was selected as a Förster resonance energy transfer (FRET) acceptor for the QD as a donor (data not shown). Little or no FRET was observed in the PAG electrophoresis run, although the samples had significant FRET when measured in bulk solution. This result raised the question of whether QD-[peptide]N and QD-[SPA]N bioconjugates migrated at slower rates because of increases in size with increases in N, or if the differences in migration were actually from delayed starts caused by a need to strip the conjugated biomolecules from the QDs in order to squeeze through the gel pores. The larger the value of N, the longer it would take to strip off all of the protein or peptide, such that different migration times would be observed even though the stripped QDs would have the same electrophoretic mobility. This possibility was discounted primarily on the basis of the peak PL emission wavelength shift across the electropherogram peaks for the peptide conjugates. The dissociation rate of bound peptide or protein from the QD should not be dependent on the size of the QD core, and thus a trend in peak PL emission wavelength would not be expected. It is also unlikely that GSH-QD “conjugates” with N = 0 peptides were exclusively mobile in the gel. Significant tailing in the electropherogram peaks would be anticipated if this were the case but was not observed. It is therefore likely that further optimization of the PAG and experimental conditions will be needed to clearly detect FRET in QD-[peptide(dye)]N conjugates.

Optimization of gel chemistry should likely aim to increase the mobility of QDs, whether to enable faster analyses, to narrow electropherogram peak widths, or to permit resolution of QD-[protein]N conjugates with larger values of N. To this end, possible changes in PAG chemistry include the use of lower crosslinker densities (although our PAGs were already near the limit of gelation with acrylamide and bisacrylamide), the substitution of bisacrylamide with longer length crosslinkers such as poly(ethylene glycol) bis-methacrylate, or the use of a porogen. Non-poly(acrylamide) hydrogel chemistries that support larger pore sizes may also be useful for establishing an optimized “stubby” capillary gel electrophoresis technique for the high-throughput characterization of QDs and their bioconjugates.

PAGE versus CE

Another useful comparison for our “stubby” capillary PAGE is conventional capillary electrophoresis (CE) where the capillary has long length and small inner diameter. The most common implementation of CE with QD-bioconjugates has been capillary zone electrophoresis (CZE) with only background electrolyte,45,46 although there have been examples of the use of polymer sieving solutions for separation of QDs.47–49

CE has offered analysis times that range from approximately 25–50% of those with our “stubby” PAG-filled capillary, albeit at applied voltages that are 1–2 orders of magnitude larger. CE has been used to demonstrate the separation of different sizes of QDs,48–50 and to confirm conjugation with peptides and proteins.51–55 Many of these analyses have relied on differences in migration time with one-colour detection of PL emission,51–53,56 whereas other analyses utilized two-colour detection for FRET within a conjugate.54,55,57 To our knowledge, only one CE study has measured zeta potentials58 and, in surveying the literature, we found the resolution of peptide and protein conjugates to be similar between CE methods and our PAGE method. Most importantly, detailed spectral analyses of electropherogram bands have been overlooked and we are not aware of any CE study that has directly compared different QD nanocrystals, different coatings, and different bioconjugates.

To help fill the gaps in the current literature, we carried out preliminary experiments to compare CZE and PAGE between different sizes of QDs and different coatings. The results, which are reported in the ESI, indicated that CZE also had an ability to resolve different QD sizes and coatings, albeit via a mechanism dominated by charge rather than size. This mechanistic difference, and the resulting similarities and differences between the CZE and PAGE data, suggested that the two methods have complementary capabilities. The combination of CZE and PAGE is thus likely to provide more complete characterization than one method alone.

Other materials

Although this study has focused on the characterization of QDs, our “stubby” capillary PAGE method is potentially useful for the characterization of other similarly sized nanoparticles and their coatings. The ESI has proof-of-concept data for the characterization and resolution of gold nanoparticles (Au NPs) of two different sizes and with two different ligand coatings. Given the different optical properties of Au NPs versus QDs, detection was via optical extinction rather than emission. Moreover, spectrofluorimetric detection and native PAGE separation of fluorescent dye-labeled proteins was also possible, as shown in the ESI. The “stubby” capillary format for PAGE thus has a range of potential applications beyond the characterization of QDs.


We revisited the PAGE of QDs and their bioconjugates by utilizing a novel “stubby” capillary format with spectrofluorimetric detection. This format improved on the current state-of-the-art by enabling more rapid and more detailed characterization of QDs than was possible with both PAG and agarose slab gels. Correlations, or the lack thereof, between the peak PL emission wavelength and the electropherogram peaks provided new insight into the key factors that determine electrophoretic mobility, and helped to resolve heterogeneity and sub-populations in ensembles of QDs. Even greater insight was possible when combined with a Ferguson analysis. We demonstrated the application of PAGE to the characterization of QDs, and particularly their functionalization with ligand and polymer coatings and conjugation with peptide and protein molecules. With further development and optimization, we anticipate that capillary PAGE with spectrofluorimetric detection will become a valuable addition to the toolbox of analytical techniques for the characterization QDs and other nanoparticle materials, their coatings, and their bioconjugates.

Conflicts of interest

There are no conflicts to declare.


The authors thank the Natural Sciences and Engineering Research Council of Canada (NSERC), the Canada Foundation for Innovation (CFI), BCKDF, and the University of British Columbia for support of this research. H. K. is grateful for a 4YF support from the University of British Columbia and a postgraduate scholarship from NSERC. T. J. and M. V. T. are grateful for support from NSERC through the CREATE NanoMat training program. W. R. A. is grateful for a Canada Research Chair (Tier 2), a Michael Smith Foundation for Health Research Scholar Award, and Alfred P. Sloan Research Fellowship. The authors also thank Brian Ditchburn (glassblower), Francis Manalastas (Electronic Engineering Services), and Pritesh Padhiar (Mechanical Engineering Services) in UBC Chemistry for help with construction of the apparatus, and Ben Herring for assistance with CE.


  1. I. L. Medintz, H. T. Uyeda, E. R. Goldman and H. Mattoussi, Nat. Mater., 2005, 4, 435–446 CrossRef CAS PubMed .
  2. X. Michalet, F. F. Pinaud, L. A. Bentolila, J. M. Tasay, J. J. Li, G. Sundaresan, A. M. Wu, S. S. Gambhir and S. Weiss, Science, 2005, 307, 538–544 CrossRef CAS PubMed .
  3. U. Resch-Genger, M. Grabolle, S. Cavaliere-Jaricot, R. Nitschke and T. Nann, Nat. Methods, 2008, 5, 763–775 CrossRef CAS PubMed .
  4. N. Hildebrandt, C. M. Spillmann, W. R. Algar, T. Pons, M. H. Stewart, E. Oh, K. Susumu, S. A. Diaz, J. B. Delehanty and I. L. Medintz, Chem. Rev., 2017, 117, 536–711 CrossRef CAS PubMed .
  5. H. Mattoussi, G. Palui and H. B. Na, Adv. Drug Delivery Rev., 2012, 64, 138–166 CrossRef CAS PubMed .
  6. E. Petryayeva, W. R. Algar and I. L. Medintz, Appl. Spectrosc., 2013, 67, 215–252 CrossRef CAS PubMed .
  7. S. J. Rosenthal, J. C. Chang, O. Kovtun, J. R. McBride and I. D. Tomlinson, Chem. Biol., 2011, 18, 10–24 CrossRef CAS PubMed .
  8. K. D. Wegner and N. Hildebrandt, Chem. Soc. Rev., 2015, 44, 4792–4834 RSC .
  9. J. B. Blanco-Canosa, M. Wu, K. Susumu, E. Petryayeva, T. L. Jennings, P. E. Dawson, W. R. Algar and I. L. Medintz, Coord. Chem. Rev., 2014, 263–264, 101–137 CrossRef CAS .
  10. K. E. Sapsford, K. M. Tyner, B. J. Dair, J. R. Deschamps and I. L. Medintz, Anal. Chem., 2011, 83, 4453–4488 CrossRef CAS PubMed .
  11. I. L. Medintz, T. Pons, S. A. Trammell, A. F. Grimes, D. S. English, J. B. Blanco-Canosa, P. E. Dawson and H. Mattoussi, J. Am. Chem. Soc., 2008, 130, 16745–16756 CrossRef CAS PubMed .
  12. C. Zhang, T. N. Do, X. W. Ong, Y. T. Chan and H. S. Tan, Chem. Phys., 2016, 481, 157–164 CrossRef CAS .
  13. J. McBride, J. Treadway, L. C. Feldman, S. J. Pennycook and S. J. Rosenthal, Nano Lett., 2006, 6, 1496–1501 CrossRef CAS PubMed .
  14. S. H. Tolbert and A. P. Alivisatos, J. Chem. Phys., 1995, 102, 4642–4656 CrossRef CAS .
  15. J. Jasieniak, L. Smith, J. van Emden, P. Mulvaney and M. Califano, J. Phys. Chem. C, 2009, 113, 19468–19474 CAS .
  16. W. W. Yu and X. Peng, Chem. Mater., 2003, 15, 2854–2860 CrossRef CAS .
  17. F. Morgner, D. Geißler, S. Stufler, N. G. Butlin, H. G. Löhmannsröben and N. Hildebrandt, Angew. Chem., Int. Ed., 2010, 49, 7570–7574 CrossRef CAS PubMed .
  18. K. D. Wegner, F. Morgner, E. Oh, R. Goswami, K. Susumu, M. H. Stewart, I. L. Medintz and N. Hildebrandt, Chem. Mater., 2014, 26, 4299–4312 CrossRef CAS .
  19. Y. X. Yang, Y. Zheng, W. R. Cao, A. Titov, J. Hyvonen, J. R. Manders, J. G. Xue, P. H. Holloway and L. Qian, Nat. Photonics, 2015, 9, 259–266 CrossRef CAS .
  20. Z. Fang, Y. Li, H. Zhang, X. H. Zhong and L. Y. Zhu, J. Phys. Chem. C, 2009, 113, 14145–14150 CAS .
  21. I. Moreels, K. Lambert, D. De Muynck, F. Vanhaecke, D. Poelman, J. C. Martins, G. Allan and Z. Hens, Chem. Mater., 2007, 19, 6101–6106 CrossRef CAS .
  22. B. T. Zhang, R. Hu, Y. C. Wang, C. B. Yang, X. Liu and K. T. Yong, RSC Adv., 2014, 4, 13805–13816 RSC .
  23. J. K. Cooper, A. M. Franco, S. Gul, C. Corrado and J. Z. Zhang, Langmuir, 2011, 27, 8486–8493 CrossRef CAS PubMed .
  24. T. Pons, H. T. Uyeda, I. L. Medintz and H. Mattoussi, J. Phys. Chem. B, 2006, 110, 20308–20316 CrossRef CAS PubMed .
  25. E. E. Lees, M. J. Gunzburg, T. L. Nguyen, G. J. Howlett, J. Rothacker, E. C. Nice, A. H. A. Clayton and P. Mulvaney, Nano Lett., 2008, 8, 2883–2890 CrossRef CAS PubMed .
  26. R. Westermeier, Electrophoresis in Practice: A Guide to Methods and Applications of DNA and Protein Separations, Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim, 4th edn, 2005 Search PubMed .
  27. D. E. Prasuhn, J. R. Deschamps, K. Susumu, M. H. Stewart, K. Boeneman, J. B. Blanco-Canosa, P. E. Dawson and I. L. Medintz, Small, 2010, 6, 555–564 CrossRef CAS PubMed .
  28. K. Susumu, E. Oh, J. B. Delehanty, J. B. Blanco-Canosa, B. J. Johnson, V. Jain, W. J. Hervey, W. R. Algar, K. Boeneman, P. E. Dawson and I. L. Medintz, J. Am. Chem. Soc., 2011, 133, 9480–9496 CrossRef CAS PubMed .
  29. T. C. Liu, J. H. Wang, H. Q. Wang, H. L. Zhang, Z. H. Zhang, X. F. Hua, Y. C. Cao, Y. D. Zhao and Q. M. Luo, J. Biomed. Mater. Res., Part A, 2007, 83, 1209–1216 CrossRef PubMed .
  30. H. L. Zhang, Y. Q. Li, J. H. Wang, X. N. Li, S. Lin, Y. D. Zhao and Q. M. Luo, J. Biomed. Opt., 2010, 15, 015001 CrossRef PubMed .
  31. H. Kurt, M. Yüce, B. Hussain and H. Budak, Biosens. Bioelectron., 2016, 81, 280–286 CrossRef CAS PubMed .
  32. H. Y. Li and T. Q. Vu, Nano Lett., 2007, 7, 1044–1049 CrossRef PubMed .
  33. J. J. Li, Y. A. Wang, W. Z. Guo, J. C. Keay, T. D. Mishima, M. B. Johnson and X. G. Peng, J. Am. Chem. Soc., 2003, 125, 12567–12575 CrossRef CAS PubMed .
  34. W. W. Yu and X. Peng, Angew. Chem., Int. Ed., 2002, 41, 2368–2371 CrossRef CAS PubMed .
  35. K. E. Sapsford, T. Pons, I. L. Medintz, S. Higashiya, F. M. Brunel, P. E. Dawson and H. Mattoussi, J. Phys. Chem. C, 2007, 111, 11528–11538 CAS .
  36. F. Aldeek, M. Safi, N. Q. Zhan, G. Palui and H. Mattoussi, ACS Nano, 2013, 7, 10197–10210 CrossRef CAS PubMed .
  37. S. Park, N. Sinha and K. Hamad-Schifferli, Langmuir, 2010, 26, 13071–13075 CrossRef CAS PubMed .
  38. K. A. Ferguson, Metabolism, 1964, 13, 985–1002 CrossRef CAS .
  39. R. E. Anderson and W. C. W. Chan, ACS Nano, 2008, 2, 1341–1352 CrossRef CAS PubMed .
  40. M. Wu, M. Massey, E. Petryayeva and W. R. Algar, J. Phys. Chem. C, 2015, 119, 26183–26195 CAS .
  41. H. H. Girault, Analytical and Physical Electrochemistry, Marcel Dekker, New York, 2004 Search PubMed .
  42. T. Pons, I. L. Medintz, X. Wang, D. S. English and H. Mattoussi, J. Am. Chem. Soc., 2006, 128, 15324–15331 CrossRef CAS PubMed .
  43. R. S. Dubrow, in Capillary Electrophoresis: Theory & Practice, ed. P. D. Grossman and J. C. Colburn, Academic Press, San Diego, 1992, p. 141 Search PubMed .
  44. C. H. Lee, A. J. Crosby, T. Emrick and R. C. Hayward, Macromol., 2014, 47, 741–749 CrossRef CAS .
  45. F. Sang, X. Huang and J. Ren, Electrophoresis, 2014, 35, 793–803 CrossRef CAS PubMed .
  46. M. Stanisavljevic, M. Vaculovicova, R. Kizek and V. Adam, Electrophoresis, 2014, 35, 1929–1937 CrossRef CAS PubMed .
  47. G. Vicente and L. A. Colón, Anal. Chem., 2008, 80, 1988–1994 CrossRef CAS PubMed .
  48. Y. Q. Li, H. Q. Wang, J. H. Wang, L. Y. Guan, B. F. Liu, Y. D. Zhao and H. Chen, Anal. Chim. Acta, 2009, 647, 219–225 CrossRef CAS PubMed .
  49. X. Song, L. Li, H. Qian and N. Fang, Electrophoresis, 2006, 27, 1341–1346 CrossRef CAS PubMed .
  50. C. Carrillo-Carrión, Y. Moliner-Martínez, B. M. Simonet and M. Valcárcel, Anal. Chem., 2011, 83, 2807–2813 CrossRef PubMed .
  51. X. Huang, J. Weng, F. Sang, X. Song, C. Cao and J. Ren, J. Chromatogr. A, 2006, 1113, 251–254 CrossRef CAS PubMed .
  52. J. Wang, X. Huang, F. Zan, C. Guo, C. Cao and J. Ren, Electrophoresis, 2012, 33, 1987–1995 CrossRef CAS PubMed .
  53. J. Wang, J. Li, J. Li, Y. Qin, C. Wang, L. Qiu and P. Jiang, Electrophoresis, 2015, 36, 1523–1528 CrossRef CAS PubMed .
  54. J. Wang and J. Xia, Anal. Chim. Acta, 2012, 709, 120–127 CrossRef CAS PubMed .
  55. J. Wang, J. Fan, J. Li, L. Liu, J. Wang, P. Jiang, X. Liu and L. Qiu, J. Sep. Sci., 2017, 40, 933–939 CrossRef CAS PubMed .
  56. M. Stanisavljevic, J. Chomoucka, S. Dostalova, S. Krizkova, M. Vaculovicova, V. Adam and R. Kizek, Electrophoresis, 2014, 35, 2587–2592 CrossRef CAS PubMed .
  57. Y. Q. Li, J. H. Wang, H. L. Zhang, J. Yang, L. Y. Guan, H. Chen, Q. M. Luo and Y. D. Zhao, Biosens. Bioelectron., 2010, 25, 1283–1289 CrossRef CAS PubMed .
  58. I. Vorácová, K. Kelpárník, M. Lisková and F. Foret, Electrophoresis, 2015, 36, 867–874 CrossRef PubMed .


Electronic supplementary information (ESI) available: Methods for coating QDs with GSH, DHLA, DHLA-SB, API-PMA, and HDA-PMA; synthesis of DHLA-SB, HDA-PMA and API-PMA; TEM images of QDs; absorbance and PL emission measurements for QDs in bulk solution; further details on the analysis of electropherogram peaks and Ferguson analysis; additional electropherograms and PL spectra; details on the theoretical correlation between peak PL emission wavelength, migration time, and QD core size; PAGE data with Au NPs and dye-labeled proteins; CZE data with QDs. See DOI: 10.1039/c7an01581j

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