Pore-Scale Investigation of Nanoparticle Transport in SaturatedPorous Media Using Laser Scanning CytometryRyan May,† Simin Akbariyeh,‡ and Yusong Li*,‡

†Mechanical Sales, Inc., 7222 South 142nd Street, Omaha, Nebraska 68138, United States‡Department of Civil Engineering, University of Nebraska-Lincoln, 2200 Vine Street, Lincoln, Nebraska 68583, United States

*S Supporting Information

ABSTRACT: Knowledge of nanoparticle transport and retentionmechanisms is essential for both the risk assessment and environmentalapplication of engineered nanomaterials. Laser scanning cytometry, anemerging technology, was used for the first time to investigate thetransport of fluorescent nanoparticles in a microfluidic flow cell packedwith glass beads. The laser scanning cytometer (LSC) was able toprovide the spatial distribution of 64 nm fluorescent nanoparticlesattached in a domain of 12 mm long and 5 mm wide. After 40 pV ofinjection at a lower ionic strength condition (3 mM NaCl, pH 7.0),fewer fluorescent nanoparticles were attached to the center of the flowcell, where the pore-scale velocity is relatively higher. After a longer injection period (300 PV), more were attached to the centerof the flow cell, and particles were attached to both the upstream and downstream sides of a glass bead. Nanoparticles attachedunder a higher ionic strength condition (100 mM NaCl, pH 7.0) were found to be mobilized when flushed with DI water. Themobilized particles were later reattached to some favorable sites. The attachment efficiency factor was found to reduce with anincrease in flow velocity. However, torque analysis based on the secondary energy minimum could not explain the observedhydrodynamic effect on the attachment efficiency factor.

1. INTRODUCTION

The recent revolution of nanotechnology has brought morethan 1300 nanotechnology-enabled consumer products into themarketplace.1,2 Strong evidence3−7 has suggested that nano-materials may enter the subsurface environment via directleaking from underground waste sites, reuse of wastewater, andagricultural use of biosolids containing engineered nano-materials. The concerns and also demonstrated evidencesuggested that some engineered nanomaterials may imposenegative impacts to the environment and human health.Understanding the fate and transport of nanomaterials in thesubsurface porous media will provide critical information forrisk assessment and regulation policy development.8

Nanoparticles are defined as a subclass of colloid particlesthat have at least one dimension less than 100 nm. Traditionalcolumn experiments were conducted to investigate thetransport and retention of different types of nanomaterials(e.g., nano zerovalent iron,9 fullerene C60,

10−12 carbonnanotubes,13 and nano titanium dioxide14) in porous media.Typical data from a column experiment include (i) break-through curves representing averaged aqueous concentration ofparticles in the column effluent over time and (ii) retentionprofiles representing averaged attached phase concentrationalong the column. Although these two pieces of informationcan provide valuable insights, they do not clearly distinguish theways that spatial and temporal changes to the hydrodynamicconditions affect particle transport.

To better reveal the mechanisms controlling colloidtransport, various noninvasive visualization techniques, suchas light transmission technique,15 UV−fluorescence tech-nique,16 magnetic resonance imaging,17 and X-ray tomog-raphy18 were developed. While those efforts provided improvedinsights, they targeted at a continuum scale, so that theinfluence of pore-scale hydrodynamics and porous mediumgeometry on particle deposition are not directly revealed. Inrecent years, pore-scale experimental techniques have been verysuccessful in directly observing and visualizing the transportand retention of colloidal particles.19−21 Such techniquesgenerally require very high resolution to observe particles atthe nanoscale. Typically, only several pore spaces can beobserved if particles are smaller than one micrometer.Observation at a larger scale is necessary to quantitativelysimulate nanoparticle transport in porous media.In this work, we explored the possibility of using laser

scanning cytometry to obtain the spatial distribution ofnanoscale particles in a porous medium domain at thecentimeter scale. Laser scanning cytometry is an emergingtechnology used in the biomedical field to image andquantitatively analyze individual cells in tissues in situ.22 Alaser scanning cytometer (LSC) comprises an optics unit that

Received: May 2, 2012Revised: August 6, 2012Accepted: August 25, 2012Published: August 26, 2012

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generates the laser scanning beam, an upright epifluorescencemicroscope with a motorized stage to allow for the generationof sample scan images, and a computer to acquire and analyzescan data (ComputCyte Corporation, Westwood, MA).Compared with traditional flow cytometry technology, whichfocuses on measuring particles suspended in a stream of liquid,a LSC is capable of quantifying fluorescent cells or particlesfixed on a solid substrate, which makes it possible to detectnanoparticles attached to the collector surface. Compared withlaser-based confocal microscopy, a LSC is able to scan relativelylarge sample areas without needing to refocus the instrument,which makes this technology suitable for investigating nano-particle distributions in a porous medium domain at thecentimeter scale.In this study, we demonstrated the possibility of using a LSC

as an effective tool for investigating the retention ofnanoparticles in porous media. Nanoparticles were injectedinto a microfluidic flow cell packed with glass beads. The spatialdistributions of attached nanospheres in porous media undervarious solution chemistries and flow velocities were developedbased on the LSC scanning results. The mechanismscontrolling nanoparticle transport were analyzed based on theLSC scanning results.

2. MATERIALS AND METHODS2.1. Model Particles and Porous Media. Carboxyl-

modified polystyrene latex fluorescent nanospheres (Phosphor-ex, Inc., Fall River, MA) with a manufacturer-reported meandiameter of 57 nm were used as model particles in this study.The fluorescent dye used to mark the particles was green withan excitation maximum at 480 nm and an emission maximum at520 nm. The density of the particles was 1.06 g/cm3. Theparticle-size distributions were measured using a 90Plus particlesize analyzer (Brook-haven Instruments Corporation, Holts-ville, NY) at all solution chemistry investigated.Glass beads conforming to a 20−30 U.S. mesh size (Potters

Industries, Inc., Valley Forge, PA) were selected as the modelporous medium. The glass beads were further sieved using asize 25 U.S. sieve corresponding to 710 μm, resulting inparticles with diameters ranging from 600 to 710 μm. Prior touse, the glass beads were washed to remove impurities fromtheir surfaces, following the procedure reported in Wang et al.12

The electrophoretic mobilities of the nanospheres and glassbeads were determined using a ZetaPALS analyzer (Brook-haven Instruments Corporation, Holtsville, NY). The proce-dure followed to measure the electrophoretic mobilities of theglass beads is the same as that reported by Kuznar andElimelech.23

2.2. Flow Cell Experiment. An ibidi μ-Slide I0.8 luer flowcell was purchased from ibidi LLC, Verona, WI; the apparatusconsisted of a hydrophobic plastic designed for high-resolutionmicroscopic analysis. The optical quality of the material iscomparable to that of glass and exhibits low birefringence andautofluorescence. The flow cell is 25.5 mm wide and 75.5 mmlong. A rectangular flow channel is located at the center of flowcell. The flow channel measures 5 mm in width, 50 mm inlength, and 0.8 mm in height. The inlet and outlet of the flowchannel are connected to tubes and pumps via luer adapters.The design of the flow cell is shown in Figure S1 of theSupporting Information. One layer of cleaned glass beads waspacked in the flow channel, which resulted in a porous mediumwith porosity in the range 0.45−0.49. Before each experiment,the flow cell was saturated with a desirable electrolyte injected

using a syringe pump (KD Scientific, Inc., Holliston, MA)through TYGON 3350 sanitary silicone tubing with a 1/16 in.inner diameter (Saint-Gobain Performance Plastics Corpora-tion, Beaverton, MI). A fluorescent nanosphere suspension witha solids content of 0.0025% in 3 mM or 100 mM NaCl waspumped into the flow cell at a Darcy velocity of 0.02, 0.04, 0.06,or 0.08 cm/s. After injecting 40 PV of the nanospheresuspension, 10 PV of a background solution with sameelectrolyte composition and concentration was injected.Experiments were conducted in duplicate under variousvelocity conditions.

2.3. Laser Scanning Cytometry Scanning. A LSCinstrument (CompuCyte Corporation, Westwood, MA)equipped with an Olympus BX-50 fluorescence microscopewas used to scan the nanospheres attached to the glass beads inthe flow cell. The LSC was equipped with a violet laser (405nm), 20 mW argon ion laser (488 nm, Cyonics Uniphasemodel 2014A-20SL), and a 5 mW red HeNe laser (633 nm,Cyonics Uniphase), each of which provides a single wavelengthto excite fluorescently dyed nanospheres. Four photomultipliertube (PMT) detectors are available on the LSC, each of whichmeasures a different range of wavelengths. The nanosphereswere excited with an argon ion laser (488 nm, 20 mW) anddetected through the D: 530/30 (FITC, Green FluorescentProtein) optical filter. A schematic diagram of the LSC setup isprovided in Figure S2 of the Supporting Information.The ibidi μ-Slide I0.8 luer flow cell was mounted on the

motorized stage of the fluorescence microscope. The micro-scope, operated under a 20× magnification, was slowly broughtinto focus until the top of the glass beads just come into focus.A scanning area was then determined as the middle section ofthe flow channel measuring 12 mm long and 5 mm wide. Therestriction of the scanning length occurred because the upwardprotrusion of the elbow luer adapters from the flow cell washigh enough to contact the microscope objective. The LSCscan occurred in 0.5 μm steps, and it took about 20 min tofinish scanning the defined scan area. For each scan, the LSCmeasured the emitted fluorescence of the nanospheres and alsoprecisely recorded the x and y coordinates of each recordingevent.

2.4. Interaction Energy Profile. Electrostatic interactionsand van der Waals interactions were considered whencalculating the interaction energy profile. The sphere−planeinteraction expressions developed by Guzman et al.24 using asurface element integration technique that excludes theDerjaguin approximation were used for the system ofnanospheres and glass beads in this study. The Hamakerconstant of 1 × 10−20 J was used for the glass−water−latexsystem based on previous studies25,26 of similar systems.Equations used for the interaction energy calculation areprovided in the Supporting Information.

2.5. Flow Field Simulation. Computational fluid dynamics(CFD) simulations were performed to mimic the three-dimensional (3D) flow fields in the flow cell packed withglass beads. The purpose of the CFD simulations was to revealthe influence of the pore-scale 3D fluid flow fields onnanosphere transport and attachment. A glass bead packingwas constructed for CFD simulation, based on one real flowcell experiment. The detailed description on flow cell packinggeneration and CFD simulation domain are provided in theSupporting Information. Due to the computational intensity ofthe CFD simulation, only the middle 5 mm of the flow cell wassimulated. Flow fields with Reynolds numbers (Re = ρlqd50/μ,

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where ρl is the liquid density, q is the Darcy velocity, d50 is themean grain diameter, and μ is the dynamic viscosity of thefluid) of 0.51, 0.39, 0.25, and 0.13, corresponding to Darcyvelocities of 0.08, 0.04, 0.06, and 0.02 cm/s, respectively, weresimulated. The COMSOL Multiphysics 4.2a CFD module(COMSOL, 2012) was used for these simulations.

3. RESULTS AND DISCUSSION

3.1. Electrokinetic Properties of Latex Particles andPorous Media. Dynamic light scattering measurementsrevealed that the mean diameter of the fluorescent micro-spheres changed from 64.2 to 59.8 nm when the ionic strengthincreased from 3 to 100 mM (NaCl, pH 7.0), as shown inFigure S5 of the Supporting Information. The electrophoreticmobility values of the nanospheres and glass beads as a functionof ionic strength (NaCl, pH 7.0) are shown in Figure S6 of theSupporting Information. Both the nanosphere and glass beadsurfaces were negatively charged. The electrophoretic mobilityincreased from −3.3 to −2.5 (μm/s)/(V/cm) and from −4.2 to−1.5 (μm/s)/(V/cm) for the nanospheres and glass beads,respectively, when the ionic strength increased from 3 to 100mM (NaCl, pH 7.0). Although a clear increase in theelectrophoretic mobility occurred between ionic strengths of20 and 50 mM, the electrophoretic mobility of the nanosphereswas not as sensitive to the ionic strength as that of the glassbeads, which may be attributed to the resistance of a carboxyllayer on the particle surface to the double-layer compressioneffect.27

The interaction energy profile calculations shown in FigureS7 of the Supporting Information revealed the presence of arepulsive energy barrier and a small secondary energy wellminimum at ionic strengths of 3−50 mM NaCl for thenanosphere−glass bead system. The energy barrier andsecondary minimum energy disappeared at ionic strengths of100 mM NaCl, which indicated favorable conditions for particledeposition.3.2. Flow Field in Flow Cell. Figure 1 illustrates the

simulated velocity field mimicking the flow field in the flow cellpacked with glass beads. Although the glass bead configurationin the flow cell may be different for each experiment, it is

expected that the characteristics of the flow field in thisparticular glass bead packing should represent a typical flowfield because the key parameters, such as the mean diameter ofthe glass beads, the width and depth of the flow cell, and theReynolds number of the flow field, were consistent between theexperiments and the model. The flow velocity field revealed byCFD simulation was generally nonuniform in three dimensions.As shown in Figure 1A, flow velocity close to the top of theflow cell is obviously higher than that close to the flow cellbottom, which is due to a gap between flow cell (0.8 mm high)and the top of glass beads with an average diameter of 0.64 mm.Figure 1B, C showed plane views of flow streamline andvelocity distribution at z = 0.29 and 0.59 mm, respectively.Clearly, velocity in the upper plane (Figure 1C) is generallyhigher than the velocity in the lower plane (Figure 1B), both ofwhich are highly nonuniform along the length of flow cell andin the transverse-flow direction. Several apparent preferentialflow pathways were noted by flow streamlines. In the porespaces, velocity was highest at the pore centers and vanishedalong the collector surfaces due to the nonslip boundaryconditions. It is expected that the variation of velocity field willhave significant impacts on nanoparticle transport.

3.3. Distribution of Attached Nanospheres. Represen-tative distributions of the detected fluorescence emitted fromthe attached nanospheres are shown in Figure 2. The

experiments were conducted in duplicate to ensure thereproducibility of the data. Figure S8 in the SupportingInformation provides example LSC scans for two flow-cellexperiments conducted as duplicates.Figure 2A represents an LSC scan of a 12 mm long section in

the middle of the flow cell (i.e., from 19 to 31 mm away fromthe inlet) after the injection of 40 PV nanospheres followed bythe injection of 10 PV of background solution (i.e., 3 mMNaCl, pH 7.0) at a Darcy velocity of 0.08 cm/s. The color barrepresents the amount of detected fluorescence, for which ahigher value corresponds to more nanospheres attached to thatlocation. Clearly shown here, the distribution of attached

Figure 1. (A) Domain dimension and velocity distribution for CFDsimulation at a Darcy velocity of 0.08 cm/s; (B) velocity distribution ata plane z = 0.29 mm; and (C) velocity distribution at a plane z = 0.58mm.

Figure 2. Distributions of detected fluorescence emitted from attachednanospheres at 3 mM NaCl, pH 7.0, Darcy velocity 0.08 cm/s afterinjection of (A) 40 PV and (B) 300 PV nanosphere suspensionfollowed by injection of 10 PV of background solution.

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nanospheres along the transverse direction was not uniform.Obviously, fewer nanospheres were detected in the centralregion of the flow cell. Examining the flow field simulation(Figure 1) showed several higher flow velocity regions in thedomain, which are located close to the center region of the flowcell. Correspondingly, flow velocity adjacent to the lateralboundaries of the flow cell was relatively lower due to walleffects. The flow velocity at the pore scale directly influencesthe single collector efficiency factor η0, which is a parameterreflecting the frequency of particle collisions with the grainsurface.28 Although η0 is typically considered to be influencedby the combined effects of sedimentation, interception, anddiffusion, diffusion is expected to be the controlling factor fornanoparticles. For nanospheres with an average diameter of64.2 nm (3 mM NaCl, pH 7.0), the diffusion efficiency ηDaccounts for 99.8% of η0 and has a reverse power-lawrelationship with velocity, that is, ηD ≈ v−0.75, according to acorrelation developed by Long and Hilpert.29 Therefore, it isreasonable to believe that the lower degree of attachment in thecentral part of the flow cell is due to the relatively higher localvelocity around the glass beads in the region.For the middle 12 mm section of the flow cell, the

distribution of attached nanospheres did not have an apparentchange along the flow direction, which however, does notnecessarily indicate a uniform distribution along the entirelength of flow cell. In the current experimental setup, the LSCwas not able to scan flow cell inlet due to the restriction of anupward protrusion of the elbow luer adapter (SupportingInformation, Figure S1), although previous research30,31 havereported significant amount of deposition can occur close to theinlet.3.4. Distribution of Attached Nanospheres after

Longer Injection Duration. Figure 2B presents thedistribution of the attached nanospheres after injecting 300PV of nanospheres followed by 10 PV of background solution(i.e., 3 mM NaCl, pH 7.0) at a Darcy velocity of 0.08 cm/s.Compared with the 40 PV injection experiment (Figure 2A),the nanospheres became attached to the entire surface of theglass beads, as indicated by the many closed circles detected bythe LSC. Based on the simulated flow streamline (Figure 1B,C), the nanospheres need to migrate across the streamlines andreach the rear stagnation zones of the glass beads to producethe detected circled distribution. This phenomenon furtherindicates that diffusion is a dominating factor for nanospheretransport. As shown in Figure 2B, this particular LSCexperiment was able to delineate the silhouette of packedglass beads in the scan area, allowing us to generate the glassbead packing for CFD simulations illustrated in Figure 1.Compared with the shorter injection duration (40 PV)

experiment (Figure 2A), significantly more nanospheres wereattached in the center area of flow cell after 300 PV of injection.Furthermore, more red dots were present in Figure 2A in thecenter of each glass bead, indicating more attachment in theupper hemisphere of glass beads. The locations of theseenhanced attachment events are corresponding with apreferential flow pathway as illustrated in Figure 1C, ahorizontal plane view of the flow field located in the upperhemisphere of glass beads. Higher velocity in the flowpreferential pathway will generate lower ηD as previouslydiscussed. However, after a longer injection period, thepreferential flow pathway will bring in more nanoparticlemass than other areas, leading to more attachment in thisregion.

3.5. Mobilization and Reattachment of Nanospheres.Experiments were conducted to investigate the attachment ofnanospheres under higher ionic strength (100 mM, NaCl, pH7.0). After a typical attachment experiment, 10 PV of DI waterwas sequentially pumped into the flow cell to investigate anypotential mobilization and reattachment. The LSC scans takenbefore and after the DI water elution are provided in Figure 3.

Apparently, the nanosphere distribution patterns are differentbefore and after flushing with DI water, which indicates that theattached nanospheres were mobilized when DI water wasinjected. The observed mobilization suggested that somenanoparticles were previously not attached in the primaryenergy minimum, although the calculated energy barrierdisappeared for the nanosphere−glass bead system when theionic strength reached 100 mM (Supporting Information,Figure S7). The interaction energy calculation was based on anaveraged surface potential of the glass beads. The actual surfacecharge distribution of a glass bead however very possibly wasnot homogeneous. Most solid surfaces in aqueous media areheterogeneously charged due to either the complexity of thecrystalline structure of solids or due to their complex chemicalcomposition. On the well-cleaned glass bead surfaces, surfacecharge heterogeneity is possible due to the presence ofheterogeneously distributed hydroxyl groups.32,33 Based onFigure S7 of the Supporting Information, the interaction energyprofile just transitioned from repulsive to attractive when theionic strength increased to 100 mM. Sensitivity analysis showedthat a slight decrease (5%) of glass bead ζ potential value usedfor the calculation will lead to a low energy barrier (0.54 kT at1.15 nm) and a secondary energy minimum (−1.74 kT at 2.62nm). Thus, some nanospheres may be just loosely associatedwith the glass bead surface at areas slightly more negativelycharged, so that change of solution to DI water will lead to theirmobilization.The total amount of fluorescence detected after flushing with

DI water was approximately 1.36 times that detected beforeflushing with DI water. Given the fact that the scan area is

Figure 3. Distributions of attached nanospheres (A) before and (B)after 10 PV of DI was pumped into the flow cell following a typicalattachment experiment, where 40 PV of nanospheres was injectedfollowed by the injection of 10 PV of 100 mM NaCl at a Darcyvelocity of 0.04 cm/s.

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located in the middle along the length of flow cell, it is possiblethat some nanospheres mobilized in the upstream of the flowcell may reattach in this area, resulting in increasedfluorescence. Furthermore, the nanosphere distribution afterflushing with DI water (Figure 3B) showed much morescattered red spots compared with the distribution observedbefore DI water flushing (Figure 3A). The red spotscorresponded to greater amounts of detected fluorescence atthe given locations, indicating that multiple nanospheres weredetected in those locations, which was favorable under DI waterflushing. The observed preferential deposition onto certainfavorable spots likely reflected the influence of surface chargeheterogeneity.34−36 Although the glass beads used in theexperiments were cleaned by acids, it is extremely difficult tocompletely rule out the contributions from impurities (e.g.,metal oxide). Previous work36 has shown that the amount oflatex colloidal particle deposition onto glass beads was sensitiveto the glass bead clean methods. Typically, surface chargeheterogeneity was considered to be negligible at high pHconditions (e.g., pH 10).37 In our experiments (pH 7), it ishighly possible that nanoscale metal oxide impurity on the glassbead surfaces will form local favorable spots, attracting themobilized nanospheres.3.6. Influence of Darcy Velocity. The distributions of

nanospheres at Darcy velocities of 0.02, 0.04, and 0.06 cm/s arepresented in Figure 4. As in the experiment conducted at 0.08cm/s (Figure 2A), the attached nanosphere distributions aregenerally uniform along the longitudinal direction, but fewerare attached to the center than close to the lateral boundaries. Itseems that more full circles were formed around the glass beadsurfaces at lower velocities and that more scattered red spotswere observed at higher velocities. This observation could also

be attributed to the diffusion-control mechanism. The pore-scale velocity is generally lower at a lower Darcy velocity, sothat nanospheres diffuse more easily across the flow streamlinesand reach every point on the surfaces of the glass beads.However, a relatively higher Darcy velocity mitigates this effect.Recent studies38−41 have reported that hydrodynamic forces

can influence not only the transport of a particle to the vicinityof a collector but also the particle attachment. To evaluate thiseffect in the studied system, we estimated the attachmentefficiency factor α based on LSC scanning at different flowvelocities. By definition, α reflects the fraction of collisions thatlead to successful attachment. Under favorable conditions, all ofthe collisions lead to attachment, such that α equals 1.Assuming a linear relationship between fluorescent signal andattached nanoparticle concentration, α can be estimated as theratio of the detected amount of fluorescence betweenunfavorable I and favorable conditions If corrected by flowvelocity:

α = II

uuf

f

(1)

where u and uf are the Darcy velocities for the unfavorable andfavorable experiments, respectively. Favorable attachmentconditions were achieved using 100 mM NaCl as thebackground solution (Figure 3A), which has no energy barrieraccording to the interaction energy calculation (SupportingInformation, Figure S2). Our calculation showed that αgradually increased from 0.31 to 0.41 when the flow velocitydecreased from 0.08 to 0.04 cm/s; α then sharply increased toclose to 1.0 when the flow velocity reached 0.02 cm/s(Supporting Information, Figure S9). Previous studies havealso reported a decrease in the attachment rate with an increaseof flow rate38,41,42 under unfavorable conditions. In thesestudies, the influences of hydrodynamic forces on particleattachment were incorporated using a torque analysis approach.Successful attachment was considered to occur when theadhesive torque acting on the attached particles was greaterthan the hydrodynamic torque.We conducted torque analysis for two glass beads adjacent to

the wall of the flow cell and two glass beads in the center of theflow cell, following the procedures by Torkzaban et al.,39 whichis detailed in the Supporting Information. Briefly, thehydrodynamic torque was calculated based on the hydro-dynamic shear obtained from the computational fluid dynamicsimulation of porous media in the flow cell. The adhesivetorque was estimated based on the secondary energy minimumwell and the corresponding separation distance. About 5.9%and 9.3% of the surfaces were found to be hydrodynamicallyfavorable for attachment (or the adhesive torque is greater thanthe hydrodynamic torque) for glass beads located in the centerof the flow cell and adjacent to the flow cell lateral boundary,respectively. The lower hydrodynamically favorable surface areaof the glass beads in the center of the flow cell is consistent withthe previous observation that fewer nanospheres were detectedin the center of the flow cell. However, the calculatedpercentage of hydrodynamically favorable surface areas (Sf) isnot sensitive to the flow velocities investigated here. Althoughthe hydrodynamic torque acting on a 64.2 nm diameternanosphere is very small, it is generally larger than the adhesivetorque (6.29e−26 N·m) based on the very small secondaryminimum energy (ca. 0.03 kT) exerted rather far away from thecollector surface (i.e., 52.2 nm). Only in the flow stagnationregions is the adhesive torque larger than the hydrodynamic

Figure 4. Distributions of nanospheres after injection of 40 PV ofnanosphere suspension followed by injection of 10 PV of 3 mM NaClbackground solution at Darcy velocities of (A) 0.02 cm/s, (B) 0.04cm/s, and (C) 0.06 cm/s.

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torque and thus hydrodynamically favorable. Although particledeposition in the secondary minimum has been widelyconsidered as a key mechanism to explain deposition underunfavorable conditions,42−44 it seems insufficient to explain theobserved declining trend in the collision efficiency factor withflow velocity. Sensitivity analysis (Supporting Information,Figure S9) indicated that, if the adhesive torque wasapproximately 10 times higher than the secondary minimumand located closer (e.g., 15 nm away) to the collector surface,the dependency of α on the velocity predicted by torqueanalysis would be similar to that observed in this study. Weagain speculated that the surface charge heterogeneity of glassbeads played an important role here. Presence of metalimpurities may lead to less negative surface potentials oncertain locations of the glass beads, which will lead to higheradhesive torques.

4. ENVIRONMENTAL IMPLICATION

In this work, we demonstrated the capability of a LSC toinvestigate nanosphere deposition and mobilization at porescale. Most importantly, the LSC provided detailed distribu-tions of attached nanoparticles in a pore space in the scale ofseveral centimeters, which allows directly linking porousmedium configuration and pore-scale hydrodynamics withnanoparticle deposition. A LSC scan revealed nanospheremobilization and reattachment under DI water flushingfollowing a deposition experiment at higher ionic strength.The column breakthrough curve cannot reflect the mobilizationprocess discovered here, because most mobilized nanoparticleswere reattached when moving to the subsequent part of thecolumn, leading to minimal influence on the nanoparticleconcentration in the column effluent. The similar comparisonof retention profile before and after DI water flushing is notpossible using a column experiment, because the retentionprofile of a column experiment is typically obtained bydestructively measuring the amount attached in a differentsection of the column. The LSC served as a novel technique toallow the investigation of these phenomena for particles at thenanoscale.This work only provided proof-of-concept by using

fluorescent polystyrene nanoparticles under simple solutionchemistry and pore structures. It can be easily extended tostudy more realistic nanomaterials with natural or artificialfluorescent dyes. Further investigation on quantitatively linkingthe detected fluorescence with attached nanoparticle concen-tration is warranted.

■ ASSOCIATED CONTENT

*S Supporting InformationAdditional experimental details, interaction energy calculations,figures, and tables. This material is available free of charge viathe Internet at http://pubs.acs.org.

■ AUTHOR INFORMATION

Corresponding Author*Phone: (402) 472-5972; fax: (402) 472-8934; e-mail: yli7@unl.edu.

NotesThe authors declare no competing financial interest.

■ ACKNOWLEDGMENTSThis work was supported by the National Science Foundationunder grant CBET 1133528. We thank the Biomechanics,Biomaterials and Biomedicine (BM3) Instrumentation Facilityat the University of Nebraska-Lincoln for providing access tothe laser scanning cytometer.

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