APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Nov. 2010, p. 7292–7298 Vol. 76, No. 210099-2240/10/$12.00 doi:10.1128/AEM.00324-10Copyright © 2010, American Society for Microbiology. All Rights Reserved.

Dispersion of TiO2 Nanoparticle Agglomerates byPseudomonas aeruginosa�†

Allison M. Horst,1* Andrea C. Neal,1 Randall E. Mielke,1 Patrick R. Sislian,2 Won Hyuk Suh,3‡Lutz Madler,4,5 Galen D. Stucky,3 and Patricia A. Holden1

Donald Bren School of Environmental Science and Management, University of California at Santa Barbara, Santa Barbara,California 931061; Department of Chemical and Biomolecular Engineering, University of California at Los Angeles,

Los Angeles, California 900952; Department of Chemistry and Biochemistry, University of California atSanta Barbara, Santa Barbara, California 931063; IWT Foundation Institute of Materials Science,

Department of Production Engineering, University of Bremen, Bremen 28359, Germany4; andCalifornia NanoSystems Institute, University of California at

Los Angeles, Los Angeles, California 900955

Received 5 February 2010/Accepted 7 September 2010

Engineered nanoparticles are increasingly incorporated into consumer products and are emerging aspotential environmental contaminants. Upon environmental release, nanoparticles could inhibit bacterialprocesses, as evidenced by laboratory studies. Less is known regarding bacterial alteration of nanoparticles,including whether bacteria affect physical agglomeration states controlling nanoparticle settling and bioavail-ability. Here, the effects of an environmental strain of Pseudomonas aeruginosa on TiO2 nanoparticle agglom-erates formed in aqueous media are described. Environmental scanning electron microscopy and cryogenicscanning electron microscopy visually demonstrated bacterial dispersion of large agglomerates formed in cellculture medium and in marsh water. For experiments in cell culture medium, quantitative image analysisverified that the degrees of conversion of large agglomerates into small nanoparticle-cell combinations weresimilar for 12-h-growth and short-term cell contact experiments. Dispersion in cell growth medium was furthercharacterized by size fractionation: for agglomerated TiO2 suspensions in the absence of cells, 81% by mass wasretained on a 5-�m-pore-size filter, compared to only 24% retained for biotic treatments. Filtrate cell andagglomerate sizes were characterized by dynamic light scattering, revealing that the average bacterial cell sizeincreased from 1.4 �m to 1.9 �m because of nano-TiO2 biosorption. High-magnification scanning electronmicrographs showed that P. aeruginosa dispersed TiO2 agglomerates by preferential biosorption of nanopar-ticles onto cell surfaces. These results suggest a novel role for bacteria in the environmental transport ofengineered nanoparticles, i.e., growth-independent, bacterially mediated size and mass alterations of TiO2nanoparticle agglomerates.

The large-scale production and use of nanoparticulate tita-nium dioxide (TiO2) in a suite of consumer products (www.nanotechproject.org) make it likely that these nanoparticleswill enter the environment via waste or erosional processes (15,27). As with other potential pollutants, interactions of nano-particles with environmental bacteria could result in eitherdirect toxicity to bacteria or biotransformations which wouldinfluence nanoparticle mobility in soil and water. Studies of theeffects of bacteria on nanoparticles have mainly focused onbacterial coalescence of already-dispersed nanoparticles. Forexample, Limbach et al. (21) observed that nanoparticulateCeO2 was initially dispersed in wastewater but then sorbed toactivated sludge bacteria. Similarly, Kiser et al. (18) describedbiosorption of nanoscale TiO2 to activated sludge within awastewater treatment plant and in laboratory batch reactors.

Nanoparticles were initially dispersed and then became asso-ciated with bacterial cells or exopolymers, which would tend todecrease the mobility of individual nanoparticles.

Nanoparticle adhesion to bacterial surfaces has been ob-served in toxicity studies for a range of nanoparticle chemis-tries and bacterial species. For example, cytotoxicity of nano-CeO2 to Escherichia coli depended on nanoparticle adhesionto the bacterial outer membrane (37). Nanoparticulatefullerene (19), ZnO (10), and CdTe quantum dot (22) toxicityto bacteria was predicated upon nanoparticle-cell contact.Nanoparticle adhesion to bacteria may be by nonspecific elec-trostatic interactions (as for CeO2 [37]) or may involve otherinteractions with bacterial surface polymers. Jucker et al. (12)demonstrated irreversible adhesion of outer membrane poly-saccharides from E. coli to nanoscale TiO2 by formation ofhydrogen bonds. The affinities of nanoparticles for bacterialmembranes imply a general consequence beyond direct bacte-rial toxicity: cell biosorption of agglomerated or dispersednanoparticles will strongly influence nanoparticle mobility inthe environment.

Due to high surface area and interfacial energies (11), TiO2

nanoparticles will agglomerate in aqueous environments. Ag-glomeration driven by electrostatics promotes settling (31) andcan thus remove nanoparticles from the water column. Given

* Corresponding author. Mailing address: 2308 Bren Hall, Univer-sity of California at Santa Barbara, Santa Barbara, CA 93106-5131.Phone: (805) 893-5652. Fax: (805) 893-7612. E-mail: ahorst@umail.ucsb.edu.

‡ Present address: 306 Stanley Hall, Department of Bioengineering,University of California at Berkeley, Berkeley, CA 94720.

† Supplemental material for this article may be found at http://aem.asm.org/.

� Published ahead of print on 17 September 2010.

7292

on Novem

ber 17, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

the high affinity between many nanoparticles and bacteria,initially agglomerated nanoparticulate TiO2, upon simple con-tact with bacteria, should tend toward dispersion due to par-ticles preferentially sorbing onto cells. Here, a combination ofhigh-resolution imaging, quantitative image analysis, and size-based agglomerate fractionation is used to describe bacteriallymediated dispersion of TiO2 nanoparticle agglomerates. Thisphenomenon is likely nonspecific to either Pseudomonasaeruginosa or TiO2 nanoparticles and thus reveals a generaliz-able role for bacteria in enhancing the mobility of nanopar-ticles in the environment.

MATERIALS AND METHODS

Bacteria, nanoparticles, and other chemicals. The environmental isolate P.aeruginosa PG201 (9, 32) was used. Nanoparticulate TiO2 was industrial P25Aeroxide (Evonik, Parsippany, NJ) (75% anatase and 25% rutile) (29). To testthe dependence of some findings on nanoparticle size and morphology, labora-tory-synthesized TiO2 (23, 24) (80% anatase and 20% rutile) was also studied incell growth experiments (see the supplemental material for synthesis details).Hereinafter, P25 Aeroxide TiO2 is called “industrial” TiO2. Nanoparticles weremaintained in the dark at room temperature prior to experiments. Experimentswere performed either in Luria-Bertani (LB) broth (Fisher Scientific, Pittsburgh,PA) prepared with nanopure water (pH 6.9, 18.2 M�-cm) or in marine coastalwater (pH 7.7, conductivity, 51.5 mS) collected from the Carpinteria Salt Marshin Carpinteria, CA (34°24�N, 119°31�W) (3). All chemicals were reagent grade orbetter (Sigma Chemical or Fisher Scientific).

Zeta potential. Zeta potentials of bacterial cells and TiO2 nanoparticles weremeasured in LB media at 25°C using a Nano-Series Zeta Sizer (Nano-ZSZEN3600; Malvern Instruments, Worcestershire, United Kingdom) (see thesupplemental material).

Bacterial growth and contact experiments. Three types of experiments wereperformed under dark conditions: one where P. aeruginosa was grown for 12 hwith preagglomerated industrial TiO2 (cell growth experiment), another wherepreagglomerated industrial TiO2 was exposed for 10 min to washed, exponential-phase P. aeruginosa cells (cell contact experiment), and the last, where pre-agglomerated industrial TiO2 was added to cell-free 12-h culture supernatant(supernatant contact experiment). For cell growth experiments only, bacterialeffects on laboratory-synthesized TiO2 were also tested. Here, “preagglomer-ated” means that nanoparticles were agglomerated in sterile LB media (10 min)before bacterial inoculation. P. aeruginosa cells were grown from frozen (�80°C)stock on LB agar plates for 8 h, and then colonies were subcultured in LB media(10 ml, 200 rpm, 37°C) for 8 h; cells were determined to be mid-exponentialphase at this time (data not shown). For cell growth experiments, LB medium (10ml) containing 0.5 mg ml�1 of industrial TiO2 was inoculated with P. aeruginosato an initial optical density at 600 nm (OD600) of 0.01. Based on prior research(1), 0.5 mg ml�1 TiO2 was not expected to inhibit bacterial growth under darkconditions but did promote extensive nanoparticle agglomeration. Several 400-�laliquots were removed within 5 min of inoculation and again at late exponentialphase (12 h, 200 rpm, 37°C) for imaging by scanning electron microscopy.

For cell contact experiments, bacteria were grown for 12 h, then separated bycentrifugation (12,000 � g, 20 min). The supernatant was removed and reserved(4°C). The cell pellet was dispersed in 5 ml of fresh LB medium, followed by 5ml of 1.0-mg ml�1 industrial TiO2 in LB medium. For supernatant contactexperiments, industrial TiO2 nanoparticles were added to the culture superna-tant at 0.5 mg ml�1 and vortexed briefly. For cell contact and supernatant contactexperiments, aliquots (400 �l each) were removed within 30 min for imaging.

To test whether dispersion may also occur in natural waters, contact experi-ments were performed in filter (0.2 �m)-sterilized marsh water. As above, P.aeruginosa was grown in, and recovered from, LB broth, then washed (3 times)with nanopure water and resuspended in marsh water (5 ml). Industrial TiO2 (1.0mg ml�1) prepared in marsh water was added to the cell suspension and brieflyvortexed.

Fractionation of agglomerates by filtration. Abiotic and cell contact treat-ments were prepared in triplicate to determine the fraction of industrial TiO2

able to pass through a porous filter. Samples were syringe filtered throughmembranes (Nuclepore track-etched polycarbonate membranes, 5-�m pore size;Whatman, GE Health Care), and the filter-retained dry mass (after 8 h at 105°C)was measured. The filtrates were reserved for dynamic light scattering.

Dynamic light scattering. Agglomerate size distributions in contact experi-ment filtrates were determined by dynamic light scattering within 30 min of

filtration using the Malvern Instruments Nano-Series Zeta Sizer (as for zetapotential measurements; see above) at 25°C. Dynamic light scattering was alsoperformed for 12-h washed P. aeruginosa cells (no TiO2) treated with sodiumazide (10 min) to arrest motility, because cell swimming interfered with Brown-ian motion-based measurement of hydrodynamic diameter (not shown).

Liquid tensiometry. Since P. aeruginosa can produce surfactants during growth(34), which could contribute to dispersion, surface tension was measured as anindicator for biosurfactant production in culture supernatant. Samples weredispensed into 10 ml acid (H2SO4 plus NoChromix; Godax Laboratories, Inc.,Cabin John, MD)-washed glass beakers for measurements at room temperatureusing a K10 digital liquid tensiometer (Kruss, Hamburg, Germany) and a small-volume plate.

Transmission electron microscopy. Primary particle size and morphology forindustrial and laboratory-synthesized TiO2 nanoparticles were determined fromtransmission electron micrographs (see the supplemental material).

Environmental scanning electron microscopy. For experiments in LB broth,aliquots (400 �l) were prepared for microscopy by centrifugation (10,000 � g, 10min), washing (3 times, 400 �l nanopure H2O), and then resuspension in 100 �lnanopure H2O. Washing removed residual LB salts, which convolute images, butdid not significantly alter agglomerate size or morphology (see the supplementalmaterial). Images were acquired using an FEI Co. XL30 FEG environmentalscanning electron microscope (Philips Electron Optics, Eindoven, Netherlands)(see the supplemental material). For each treatment, 15 random images weretaken (�2,500 magnification; 5 images for each triplicate sample). For experi-ments in salt marsh water, aliquots were not washed prior to imaging and 5random images were taken for each treatment (�625 magnification).

Quantitative analysis of microscopy images. Size distributions for nanopar-ticle agglomerates in LB media were determined from environmental scanningelectron microscope images. Equivalent diameters (X) for nanoparticle agglom-erates (classified as small where 0 � X � 6 �m, medium where 6 � X � 12 �m,and large where 12 � X � 18 �m) were measured for each image using NIS-Elements Basic Research software (Nikon Instruments Inc., Melville, NY) (seethe supplemental material). Statistical analysis of size distribution data wasperformed by Tukey’s honestly significant difference test.

Cryogenic scanning electron microscopy. Specimens from growth experimentswere washed (once with nanopure H2O) prior to liquid nitrogen cryogenicfixation. Samples were transferred to a Polaron cryogenic stage (Quorum Tech-nologies, Guelph, Ontario, Canada) for etching (�70°C, 3 min), followed by goldsputter coating (3 to 5 nm). Imaging was performed using the FEI XL30 instru-ment as described above, at a 20-kV accelerating voltage and 10.3-mm workingdistance.

RESULTS

Nanoparticle and cell characteristics. Industrial TiO2 ishighly heterogeneous; thus, a size range was determined ratherthan a mean diameter. Equivalent diameters of industrial TiO2

nanoparticles (from TEM images) ranged from 6.4 nm to 73.8nm, with 75% having an equivalent diameter between 15 and60 nm. The primary particle size for laboratory-synthesizedTiO2 nanoparticles was 16.0 � 1.5 nm. The specific surfacearea of industrial TiO2 (54 m2/g) was measured by theBrunauer-Emmett-Teller (BET) technique using a Tristar3000 (Micrometrics, Norcross, GA). The specific surface areafor laboratory-synthesized TiO2 (94 m2/g) was calculated as-suming spherical particles, uniform diameter (16 nm), and anestimated density of 3.97 g/cm3 based on the weighted anatase-to-rutile ratio. The zeta potentials for P. aeruginosa and indus-trial TiO2 nanoparticles in LB media were �9.1 mV and �17.9mV, respectively.

Nanoparticle agglomerate shifts in growth experiments.TiO2 nanoparticles were highly agglomerated in LB mediaboth before and immediately after inoculation with P. aerugi-nosa (Fig. 1; see the supplemental material). Quantitative anal-ysis of environmental scanning electron microscope imagesshowed no effect of inoculation on the maximum size of TiO2

agglomerates (Table 1). The sizes of washed large agglomer-

VOL. 76, 2010 TiO2 NANOPARTICLE DISPERSION BY PSEUDOMONAS AERUGINOSA 7293

on Novem

ber 17, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

ates appeared consistent with sizes of agglomerates in un-washed LB media (see the supplemental material), suggestingthat the washing steps did not significantly alter agglomeratesize or morphology.

After 12 h under abiotic conditions, the maximum size ofindustrial TiO2 agglomerates increased while the maximumsize of laboratory-synthesized TiO2 agglomerates decreased(Table 1). In cell growth treatments, industrial TiO2 agglom-erates appeared smaller after 12 h than abiotic controls andcells and nanoparticles appeared to be highly colocalized (Fig.1B). Dispersion of large agglomerates after 12 h of bacterialgrowth was further evidenced by a 3-fold increase in the fre-quency of small agglomerates, which could be remaining abi-otic agglomerates or cells encrusted by nanoparticles, while the

frequency of large and mid-sized agglomerates was signifi-cantly decreased (Fig. 2). The maximum size of industrial TiO2

agglomerates in 12-h cultures was less than one-half of theinitial size (Table 1). At higher magnification, industrial TiO2

nanoparticle clusters were observed at individual cell surfaces(Fig. 1C). In several micrographs, bacteria appeared to beassociated with, and detaching from, large TiO2 agglomerates(Fig. 1D). Similar results were found for laboratory-synthe-sized TiO2 nanoparticles (see the supplemental material).

Dispersion of large industrial TiO2 agglomerates in P.aeruginosa cell growth experiments was also observed in cryo-genic scanning electron microscope images. Cryogenic scan-ning electron microscopy is used for visualizing nanoparticleagglomerate morphology (36), microbe-mineral interactions(4, 16), and soil colloids (28) and was used here to visuallyconfirm selected environmental scanning electron microscopyresults in light of concerns about the effect of moderate dryingand deformation, which can occur during extended imaging(17). In 12-h uninoculated samples, very large (�20-�m)highly branched TiO2 agglomerates were observed. After 12 hof cell growth, large agglomerates did not exist and bacterialcells appeared encrusted by TiO2 (Fig. 3C and E). Large ag-glomerates were also apparent at lower magnification in cryo-genic scanning electron microscope images of 12-h abioticsamples, and the specimen surface was heterogeneous andtextured. In contrast, the surfaces from 12-h cell growth treat-ments appeared smooth and homogeneous and no large ag-glomerates were observed (see the supplemental material).Thus, environmental and cryogenic scanning electron micro-scope images, in combination with quantitative data from im-

FIG. 1. Environmental scanning electron microscope images of industrial TiO2 agglomerates in uninoculated (A) and inoculated (B) LB brothafter 12 h. At higher magnification, individual cells are either circumscribed with small nanoparticle clusters (C) or are sorbed onto nanoparticleagglomerates (D). Scale bars, 10 �m (A and B), 1 �m (C), and 2 �m (D).

TABLE 1. Maximum TiO2 agglomerate sizes for cell growth, cellcontact, and supernatant contact experiments

Expt Exposure(h)

Maximum agglomerate sizea (�m) for:

IndustrialTiO2

Laboratory-synthesizedTiO2

Sterile LB media 0 12.9 � 0.8 13.4 � 1.312 16.3 � 0.8 9.8 � 0.8

Growth 0 10.8 � 1.0 13.8 � 1.512 4.9 � 0.3 5.5 � 0.4

Cell contact 0 4.9 � 0.6 NDb

Supernatant contact 0 8.7 � 0.8 ND

a Size (maximum equivalent diameter) was measured from environmentalscanning electron microscope images (n 15 images per experiment).

b ND, not determined.

7294 HORST ET AL. APPL. ENVIRON. MICROBIOL.

on Novem

ber 17, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

age analysis, demonstrate that initially large agglomerates wereextensively dispersed in P. aeruginosa growth experiments.

Contact experiments. To test if bacterial growth was re-quired for agglomerate dispersion, industrial TiO2 was broughtinto contact with either washed cells or culture supernatant, inwhich cells had been grown for 12 h and subsequently removedby centrifugation. In culture supernatant, industrial TiO2 ag-glomerates appeared smaller than agglomerates in fresh LBmedia (see the supplemental material) and the maximum ag-

glomerate size was 8.7 � 0.8 �m, which was significantlysmaller than the maximum agglomerate size in fresh LB media(Table 1). Quantitative image analysis revealed that, in com-parison to the abiotic controls (0 h), there were fewer large andmedium agglomerates and more small agglomerates in cell-free culture supernatant (Fig. 2C). However, any differences inaqueous chemistries between the fresh LB broth (Fig. 2A) andthe cell-free spent LB broth (Fig. 2C) only minimally changedthe overall agglomerate size distributions. Accumulation of

FIG. 2. Frequency of three size ranges (small [S], medium [M], and large [L]) of industrial TiO2 nanoparticle agglomerates in culture mediawith and without P. aeruginosa cells. (A and B) Experiments with uninoculated media (black bars) and inoculated media (gray bars) at 0 h (A) and12 h (B). (C) Contact experiments. Striped bars, cell-free supernatant; white bars, exponential-phase cells added to fresh LB medium. Like lettersindicate values that are not significantly different ( 0.05) for each treatment within each size range. Bars to the left of the dashed line are relatedto the left y axis; bars to the right are related to the right y axis. ESEM, environmental scanning electron microscope.

FIG. 3. Cryogenic scanning electron microscope images of industrial TiO2 nanoparticles (A), P. aeruginosa (B and D), and P. aeruginosa withindustrial TiO2 (C and E). White arrows indicate bare cells in treatments not containing TiO2 (B and D) or cells encrusted by nanoparticles (Cand E).

VOL. 76, 2010 TiO2 NANOPARTICLE DISPERSION BY PSEUDOMONAS AERUGINOSA 7295

on Novem

ber 17, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

biosurfactants did not contribute to dispersion, as the liquidsurface tensions of cell-free culture supernatant and fresh LBmedia were statistically equivalent (54.1 � 2.1 mN/m and52.3 � 2.8 mN/m, respectively).

Environmental scanning electron microscope images fromcell contact experiments revealed extensive dispersion of pre-formed industrial TiO2 nanoparticle agglomerates, whichclosely resembled the level of dispersion in 12-h cell growthexperiments (see the supplemental material). Medium andlarge agglomerates were not observed in contact experiments,and average maximum agglomerate sizes for the cell contactand cell growth experiments were statistically equivalent (4.9 �0.6 �m and 4.9 � 0.3 �m, respectively) (Table 1). Agglomeratesize distributions for the cell contact and 12-h cell growthexperiments showed a similar shift from medium and large tosmall agglomerates and cell-nanoparticle conglomerates (Fig.2). Together with scanning electron micrographs, in which cellsappear to peel away initially agglomerated nanoparticles (Fig.1), these data suggest that bacterial contact with TiO2 agglom-erates and sequestration of nanoparticle clusters onto cell sur-faces (i.e., biosorption) are the dominant processes leading todispersion.

In order to determine if bacterially mediated dispersion oc-curs under environmental conditions, contact experimentswere also performed in water collected from a salt marsh.When suspended in marsh water, TiO2 nanoparticles formedlarge agglomerates that were similar in size and morphology tothose formed in LB medium (see the supplemental material).After cell contact, large agglomerates were entirely dispersedand only a few small agglomerates existed. At higher magnifi-cation, bacteria were seen to be associated with the remainingagglomerates or separated from agglomerates and encrustedby TiO2 nanoparticles (see the supplemental material).

Agglomerate size fractionation and dynamic light scatter-ing. In abiotic samples (0 h) prepared in LB medium, 81.3% ofTiO2 by mass was retained on the filter membrane, while 24%was retained in cell contact treatments. Dynamic light scatter-ing could not be performed on the entire (unfiltered) samplesdue to the presence of medium and large (�6-�m) agglomer-ates, which interfere with the analyses. In the abiotic samplefiltrate, 51.5% of TiO2 (by volume) was agglomerated to largerthan 3 �m (Fig. 4) and 5.1% of TiO2 was in the form of verysmall (0.2- to 0.7-�m) agglomerates. This contrasted with cellcontact experiment filtrate, where 0.7% of TiO2 was agglom-erated to larger than 3 �m. The small-size (0.2- to 0.7-�m)populations in the abiotic filtrate were not observed in cellcontact filtrate (Fig. 4), suggesting that dispersed TiO2 nano-particles were removed from suspension by bacterial biosorp-tion. The average particle size in cell contact experiments(1.9 � 0.1 �m) was larger than the average size of P. aeruginosawithout TiO2 (1.4 � 0.0 �m) (Fig. 4) due to TiO2 adsorptiononto cells, as observed in scanning electron microscope images(Fig. 1C and D and 3C and E).

DISCUSSION

TiO2 nanoparticle agglomerates are held together by weakelectrostatic or van der Waals forces (38) in natural waters.The degree and stability of agglomeration depend highly onenvironmental chemistry, and abiotic factors affecting agglom-

eration are well studied. For example, divergence from thenanoparticle isoelectric point (the pH at which the net particlesurface charge is neutral) leads to interparticle electrostaticrepulsion, thereby decreasing agglomeration (7, 8). Organicacids (e.g., humic and fulvic acids) can similarly decrease ag-glomeration due to the coating of nanoparticles, leading tosteric stabilization (6). Increasing ionic strength and additionof divalent cations facilitate nanoparticle agglomeration bycompressing the electrical double layer at nanoparticle sur-faces, thereby reducing repulsion (8). Yet, while effects ofabiotic chemistry on agglomeration are well established, thepotential effects of biotic components of aqueous environ-ments, including bacteria, are not understood.

This study shows that an environmental strain of P. aerugi-nosa promotes dispersion of initially agglomerated industrialand laboratory-synthesized TiO2 nanoparticles. In cell growthexperiments, both industrial and laboratory-synthesized TiO2

nanoparticle agglomerates were largely dispersed after 12 h ofgrowth (Fig. 1; see the supplemental material). Dispersion wasdefined as an increase in the frequency of small agglomerates(Fig. 2), which could be abiotic or bacterium-nanoparticle con-glomerates, produced as a consequence of nanoparticles beingremoved from medium and large agglomerates upon biosorp-tion to bacterial cells (Fig. 1 and 3; see the supplementalmaterial). Subsequent experiments (with industrial TiO2 only)revealed that changes in culture medium chemistry, eitherfrom bacterial metabolism of medium nutrients or productionof biosurfactants, contributed little to the observed level ofdispersion. In addition to image-based methods (environ-mental and cryogenic scanning electron microscopy), filtra-tion and dynamic light scattering were used to quantify thephenomenon.

The extinction of small (0.2- to 0.7-�m) abiotic nanoparticleagglomerates in the presence of P. aeruginosa (Fig. 4) suggeststhat biosorption sequesters dispersed nanoparticles, a result

FIG. 4. Dynamic light scattering-based particle size distributionsfor filtered (�5-�m) suspensions of industrial TiO2 in uninoculatedLB media (TiO2), sodium azide-immobilized P. aeruginosa cells in theabsence of TiO2 (P. aeruginosa), and P. aeruginosa cultured with in-dustrial TiO2 (TiO2 � P. aeruginosa). The y axis indicates the percent-age of the total particle volume associated with a specific particle size.Data points represent the average volumetric percentages from tripli-cate samples (error bars represent standard errors).

7296 HORST ET AL. APPL. ENVIRON. MICROBIOL.

on Novem

ber 17, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

similar to that reported by Limbach et al. (21), where initiallydispersed CeO2 became associated with activated sludge bac-teria in wastewater. This phenomenon may be more relevantwhen considering lower concentrations of TiO2 or dilute aque-ous chemistries that are not expected to facilitate extensiveagglomeration. In this study, the high electrolyte concentrationin LB growth media (5 g liter�1 NaCl) promoted initial abioticTiO2 nanoparticle agglomeration and thus dispersion by bac-teria was observable. Amino acids in LB media (1.4 to 19.1mM) (33) also likely contributed to extensive TiO2 agglomer-ation, perhaps analogous to the agglomeration of initially dis-persed ZnS nanoparticles in the presence of cysteine (26). Yetdispersion of initially agglomerated TiO2 nanoparticles wasindependent of cell growth and solely due to cells contactingagglomerates.

In another study, positively charged CeO2 nanoparticleswere sorbed to the negatively charged surface of E. coli (37).Here, both cell and industrial TiO2 surfaces are negativelycharged in LB media (�9.05 mV and �17.88 mV, respec-tively), and thus averaged electrostatic interactions betweencells and nanoparticles would not account for their association.Cationic molecules within LB media could create cross-link-ages between negatively charged cells and nanoparticles, per-haps analogous to aggregation of positively charged gold nano-particles by citrate bridging (30). Alternatively, as generallydescribed by Jucker et al. (14), adhesion could be mediated bybacterial surface polymers. Adsorption to TiO2 has been ob-served for several bacterial cell surface polymers, includingpyoverdine, a membrane-associated siderophore producedabundantly by P. aeruginosa (25) that binds TiO2 via catecholbridges and can serve as a binding site for various metal ions(2). Cell surface lipopolysaccharides isolated from bacteriaformed hydrogen bonds with TiO2 (12), and negatively chargedhigher-order structures of bacterial lipopolysaccharide, i.e., mi-celles, sorbed onto negatively charged TiO2 (13). Li and Logan(20) reported lipopolysaccharide length-dependent adhesionof E. coli to solid surfaces: strains with relatively long lipopoly-saccharide polymers adhered more extensively to TiO2 thanstrains with truncated lipopolysaccharide chains. Lipopolysac-charides were also involved in biosorption of gold (5), silver(35), and CeO2 nanoparticles (37) to E. coli, suggesting thatlipopolysaccharides play a general role in nanoparticle adhe-sion to cell surfaces. Extracellular polymeric substances in-creased adhesion of a Pseudomonas species to TiO2 (20),though the mechanism was not reported. Sorption of TiO2

nanoparticles with P. aeruginosa in this study may similarly bedue to interactions with bacterial surface polymers, but furtherresearch in this area is needed.

In conclusion, this work presents new evidence that thepreferential adsorption of TiO2 nanoparticles to bacterial cellsurfaces can disperse nanoparticles that are agglomeratedprior to their interaction with bacteria. Bacterially mediateddispersion is not limited to LB medium, as dispersion was alsoobserved in salt marsh water samples. These findings maytherefore be transferable to natural waters, particularly thosewhich promote extensive nanoparticle agglomeration and highbacterial density such as salt- and nutrient-rich environments(e.g., estuaries and salt marshes). Bacterial involvement innanoparticle dispersion could be a generally important processfor nanoparticle mobility in the environment, but extrapolation

to other conditions is predicated upon nanoparticles agglom-erating abiotically and then associating in nanoparticle-bacte-rium combinations.

ACKNOWLEDGMENTS

Funding was provided by the U.S. Department of Energy Naturaland Accelerated Bioremediation Research program (award DE-FG02-05ER63949), the U.S. Environmental Protection Agency (EPA; STARawards R831712 and R833323), the U.S. EPA and National ScienceFoundation (under Cooperative Agreement number EF0830117), theUC Lead Campus for Nanotoxicology Training and Research Programfunded by UC TSR&TP, and the U.S. Department of Energy (grantDE-FG02-06ER64250) with support from Altair Nanotechnologies,Inc. Lutz Madler thanks the Deutsche Forschungsgemeinschaft (DFG;German Research Foundation, Forschungsstipendium MA 3333/1-1).W. H. Suh thanks the Otis Williams Postdoctoral Fellowship (SantaBarbara Fund).

BET and TEM analyses were performed at the MRL Central Fa-cilities, which are supported by the MRSEC Program of the NSF(under award no. DMR05-20415), a member of the NSF-funded Ma-terials Reserach Facilities Network.

Scanning electron microscopy was performed in the MEIAF Lab atUCSB. Any opinions, findings, and conclusions or recommendationsexpressed in this material are those of the authors and do not neces-sarily reflect the views of the National Science Foundation or theEnvironmental Protection Agency. This work has not been subjectedto EPA review, and no official endorsement should be inferred.

REFERENCES

1. Adams, L. K., D. Y. Lyon, and P. J. J. Alvarez. 2006. Comparative eco-toxicity of nanoscale TiO2, SiO2, and ZnO water suspensions. Water Res.40:3527–3532.

2. Beveridge, T. J., and R. J. Doyle (ed.). 1989. Metal ions and bacteria. Wiley& Sons, New York, NY.

3. Cao, Y. P., P. G. Green, and P. A. Holden. 2008. Microbial communitycomposition and denitrifying enzyme activities in salt marsh sediments. Appl.Environ. Microbiol. 74:7585–7595.

4. Chenu, C., and A. Jaunet. 1992. Cryoscanning electron-microscopy of mi-crobial extracellular polysaccharides and their association with minerals.Scanning 14:360–364.

5. Deplanche, K., and L. E. Macaskie. 2008. Biorecovery of gold by Escherichiacoli and Desulfovibrio desulfuricans. Biotechnol. Bioeng. 99:1055–1064.

6. Domingos, R. F., N. Tufenkji, and K. J. Wilkinson. 2009. Aggregation oftitanium dioxide nanoparticles: role of a fulvic acid. Environ. Sci. Technol.43:1282–1286.

7. Fernandez-Nieves, A., and F. J. D. Nieves. 1999. The role of zeta potential inthe colloidal stability of different TiO2/electrolyte solution interfaces. Col-loids Surf. A Physicochem. Eng. Asp. 148:231–243.

8. French, R. A., A. R. Jacobson, B. Kim, S. L. Isley, R. L. Penn, and P. C.Baveye. 2009. Influence of ionic strength, pH, and cation valence on aggre-gation kinetics of titanium dioxide nanoparticles. Environ. Sci. Technol.43:1354–1359.

9. Holden, P. A., M. G. LaMontagne, A. K. Bruce, W. G. Miller, and S. E.Lindow. 2002. Assessing the role of Pseudomonas aeruginosa surface-activegene expression in hexadecane biodegradation in sand. Appl. Environ. Mi-crobiol. 68:2509–2518.

10. Huang, Z. B., X. Zheng, D. H. Yan, G. F. Yin, X. M. Liao, Y. Q. Kang, Y. D.Yao, D. Huang, and B. Q. Hao. 2008. Toxicological effect of ZnO nanopar-ticles based on bacteria. Langmuir 24:4140–4144.

11. Israelachvili, J. 1992. Intermolecular and surface forces. Elsevier Ltd., SanDiego, CA.

12. Jucker, B. A., H. Harms, S. J. Hug, and A. J. B. Zehnder. 1997. Adsorptionof bacterial surface polysaccharides on mineral oxides is mediated by hydro-gen bonds. Colloids Surf. B Biointerfaces 9:331–343.

13. Jucker, B. A., H. Harms, and A. J. B. Zehnder. 1998. Polymer interactionsbetween five gram-negative bacteria and glass investigated using LPS mi-celles and vesicles as model systems. Colloids Surf. B Biointerfaces 11:33–45.

14. Jucker, B. A., A. J. B. Zehnder, and H. Harms. 1998. Quantification ofpolymer interactions in bacterial adhesion. Environ. Sci. Technol. 32:2909–2915.

15. Kaegi, R., A. Ulrich, B. Sinnet, R. Vonbank, A. Wichser, S. Zuleeg, H.Simmler, S. Brunner, H. Vonmont, M. Burkhardt, and M. Boller. 2008.Synthetic TiO2 nanoparticle emission from exterior facades into the aquaticenvironment. Environ. Pollut. 156:233–239.

16. Karimi-Lotfabad, S., and M. R. Gray. 2000. Characterization of contami-nated soils using confocal laser scanning microscopy and cryogenic-scanningelectron microscopy. Environ. Sci. Technol. 34:3408–3414.

VOL. 76, 2010 TiO2 NANOPARTICLE DISPERSION BY PSEUDOMONAS AERUGINOSA 7297

on Novem

ber 17, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from

17. Kirk, S., J. Skepper, and A. M. Donald. 2009. Application of environmentalscanning electron microscopy to determine biological surface structure. J.Microsc. 233:205–224.

18. Kiser, M. A., P. Westerhoff, T. Benn, Y. Wang, J. Perez-Rivera, and K.Hristovski. 2009. Titanium nanomaterial removal and release from waste-water treatment plants. Environ. Sci. Technol. 43:6757–6763.

19. Klaine, S. J., P. J. J. Alvarez, G. E. Batley, T. F. Fernandes, R. D. Handy,D. Y. Lyon, S. Mahendra, M. J. McLaughlin, and J. R. Lead. 2008. Nano-materials in the environment: behavior, fate, bioavailability, and effects.Environ. Toxicol. Chem. 27:1825–1851.

20. Li, B. K., and B. E. Logan. 2004. Bacterial adhesion to glass and metal-oxidesurfaces. Colloids Surf. B Biointerfaces 36:81–90.

21. Limbach, L. K., R. Bereiter, E. Mueller, R. Krebs, R. Gaelli, and W. J. Stark.2008. Removal of oxide nanoparticles in a model wastewater treatmentplant: influence of agglomeration and surfactants on clearing efficiency.Environ. Sci. Technol. 42:5828–5833.

22. Lu, Z. S., C. M. Li, H. F. Bao, Y. Qiao, Y. H. Toh, and X. Yang. 2008.Mechanism of antimicrobial activity of CdTe quantum dots. Langmuir 24:5445–5452.

23. Madler, L., H. K. Kammler, R. Mueller, and S. E. Pratsinis. 2002. Con-trolled synthesis of nanostructured particles by flame spray pyrolysis. J.Aerosol Sci. 33:369–389.

24. Madler, L., W. J. Stark, and S. E. Pratsinis. 2002. Flame-made ceria nano-particles. J. Mater. Res. 17:1356–1362.

25. McWhirter, M. J., A. J. McQuillan, and P. J. Bremer. 2002. Influence ofionic strength and pH on the first 60 min of Pseudomonas aeruginosa attach-ment to ZnSe and to TiO2 monitored by ATR-IR spectroscopy. ColloidsSurf. B Biointerfaces 26:365–372.

26. Moreau, J. W., P. K. Weber, M. C. Martin, B. Gilbert, I. D. Hutcheon, andJ. F. Banfield. 2007. Extracellular proteins limit the dispersal of biogenicnanoparticles. Science 316:1600–1603.

27. Mueller, N. C., and B. Nowack. 2008. Exposure modeling of engineerednanoparticles in the environment. Environ. Sci. Technol. 42:4447–4453.

28. Negre, M., P. Leone, J. Trichet, C. Defarge, V. Boero, and M. Gennari. 2004.Characterization of model soil colloids by cryo-scanning electron micros-copy. Geoderma 121:1–16.

29. Ohno, T., K. Sarukawa, K. Tokieda, and M. Matsumura. 2001. Morphologyof a TiO2 photocatalyst (Degussa, P-25) consisting of anatase and rutilecrystalline phases. J. Catalysis 203:82–86.

30. Ojea-Jimenez, I., and V. Puntes. 2009. Instability of cationic gold nanopar-ticle bioconjugates: the role of citrate ions. J. Am. Chem. Soc. 137:13320–13327.

31. Omelia, C. R. 1980. Aquasols—the behavior of small particles in aquaticsystems. Environ. Sci. Technol. 14:1052–1060.

32. Priester, J. H., P. K. Stoimenov, R. E. Mielke, S. M. Webb, C. Ehrhardt, J. P.Zhang, G. D. Stucky, and P. A. Holden. 2009. Effects of soluble cadmiumsalts versus CdSe quantum dots on the growth of planktonic Pseudomonasaeruginosa. Environ. Sci. Technol. 43:2589–2594.

33. Sezonov, G., D. Joseleau-Petit, and R. D’Ari. 2007. Escherichia coli physiol-ogy in Luria-Bertani broth. J. Bacteriol. 189:8746–8749.

34. Soberon-Chavez, G., F. Lepine, and E. Deziel. 2005. Production of rhamno-lipids by Pseudomonas aeruginosa. Appl. Microbiol. Biotechnol. 68:718–725.

35. Sondi, I., and B. Salopek-Sondi. 2004. Silver nanoparticles as antimicrobialagent: a case study on E-coli as a model for Gram-negative bacteria. J.Colloid Interface Sci. 275:177–182.

36. Tabellion, J., R. Clasen, J. Reinshagen, R. Oberacker, and M. J. Hoffmann.2002. Correlation between structure and rheological properties of suspen-sion of nanosized powders. Euro Ceramics 206:139–142.

37. Thill, A., O. Zeyons, O. Spalla, F. Chauvat, J. Rose, M. Auffan, and A. M.Flank. 2006. Cytotoxicity of CeO2 nanoparticles for Escherichia coli. Physico-chemical insight of the cytotoxicity mechanism. Environ. Sci. Technol. 40:6151–6156.

38. Tsantilis, S., and S. E. Pratsinis. 2004. Soft- and hard-agglomerate aerosolsmade at high temperatures. Langmuir 20:5933–5939.

7298 HORST ET AL. APPL. ENVIRON. MICROBIOL.

on Novem

ber 17, 2020 by guesthttp://aem

.asm.org/

Dow

nloaded from