Significance New methods for detecting and killing antibiotic-resistant, Gram-negative bacteria are of prime interest for a wide variety of applications. While phages have long been considered as potential antibacterial agents, many concerns about phage therapy stem from the fact that phages are replicating, evolvable entities whose biology is poorly understood in most cases. These concerns could be addressed by destroying the phage immediately upon use. We accomplish this by conjugating phages to gold nanorods, whose excitation by near-infrared light causes localized heating that essentially cooks nearby bacteria. Thus, the phages deliver gold nanorods to the targeted bacteria, and the nanorods destroy both bacteria and phages simultaneously. This strategy transforms phages from an evolving biological entity into a controlled, drug-like reagent.

Abstract The use of bacteriophages (phages) for antibacterial therapy is under increasing consideration to treat antimicrobial-resistant infections. Phages have evolved multiple mechanisms to target their bacterial hosts, such as high-affinity, environmentally hardy receptor-binding proteins. However, traditional phage therapy suffers from multiple challenges stemming from the use of an exponentially replicating, evolving entity whose biology is not fully characterized (e.g., potential gene transduction). To address this problem, we conjugate the phages to gold nanorods, creating a reagent that can be destroyed upon use (termed “phanorods”). Chimeric phages were engineered to attach specifically to several Gram-negative organisms, including the human pathogens Escherichia coli, Pseudomonas aeruginosa, and Vibrio cholerae, and the plant pathogen Xanthomonas campestris. The bioconjugated phanorods could selectively target and kill specific bacterial cells using photothermal ablation. Following excitation by near-infrared light, gold nanorods release energy through nonradiative decay pathways, locally generating heat that efficiently kills targeted bacterial cells. Specificity was highlighted in the context of a P. aeruginosa biofilm, in which phanorod irradiation killed bacterial cells while causing minimal damage to epithelial cells. Local temperature and viscosity measurements revealed highly localized and selective ablation of the bacteria. Irradiation of the phanorods also destroyed the phages, preventing replication and reducing potential risks of traditional phage therapy while enabling control over dosing. The phanorod strategy integrates the highly evolved targeting strategies of phages with the photothermal properties of gold nanorods, creating a well-controlled platform for systematic killing of bacterial cells.

Antibiotic-resistant bacterial infections, particularly from Gram-negative organisms, are widely recognized as an urgent threat to health worldwide (1). The development of new antibacterial agents targeting these organisms is therefore an important goal. Phages have been long proposed as antibacterial agents, and recent case studies (2, 3) and clinical trials (4) have prompted increased attention. However, treatment of infection by whole phages presents critical challenges, such as a lack of biological characterization of most phages, which may carry toxin genes or cause generalized transduction of bacterial genes (5). An interesting approach uses phages to deliver CRISPR-Cas cassettes as antimicrobials (6, 7), although this strategy faces challenges with efficient delivery to a wider range of bacterial targets (8). In addition, the pharmacokinetics and pharmacodynamics of phages are difficult to model due to their exponential replication, presenting a major barrier to clinical translation (9). Exponential replication may also lead to undesirably rapid release of bacterial endotoxins (10). A reductionist approach to avoid the problems associated with whole phages is to engineer phage-derived proteins, such as pyocins or lysins, as antibacterial agents (reviewed in ref. 11). However, some advantages of whole phages, such as avidity of the phage-displayed receptor-binding proteins (RBPs), which may increase affinity by ∼1,000× compared to recombinant RBP (12), interaction with secondary receptors on the bacterial host (13, 14), and subdiffusive search mechanisms (15, 16), may be lost. Therefore, an alternative approach is to utilize the phage for bacterial attachment, and then destroy the phages simultaneously with the bacteria, thus controlling dosage and avoiding undesired consequences while maintaining the advantages of whole phage as a delivery vehicle.

Here, we use photothermal heating as a physical mechanism that would result in both phage and host cell destruction, which can be achieved using metallic nanomaterials (17⇓⇓⇓⇓–22). These nanostructures, such as gold nanorods (AuNRs), exhibit a localized surface plasmon resonance (LSPR) upon irradiation with light, which induces coherent oscillation of the electron cloud. This energy can be released primarily as heat, leading to high local temperatures (e.g., ΔT up to ∼50 °C, depending on the laser power applied) with a half-length in the submicron range (from a single nanoparticle) to a few microns (from an ensemble of nanoparticles) (23, 24), potentially killing nearby bacterial or eukaryotic cells. The LSPR spectrum of AuNRs can be tuned by their size, allowing excitation by light in the near-infrared (NIR) “biological window” for which soft tissues are somewhat transparent. Other nanomaterials also exhibit nonspecific cytotoxic properties (e.g., nanosilver) through a variety of chemical mechanisms, but a general problem with the application of nanomaterials against bacterial infections is their lack of specificity against bacterial vs. mammalian cells, presenting a general challenge for biocompatibility (25).

To confer specificity to nanostructures, one may conjugate antibodies that target specific bacterial strains (26), following upon extensive work targeting nanoparticles for cancer cell treatment (27⇓⇓–30). However, phage-based strategies possess several advantages compared to antibody-based strategies. First, greater delivery of nanoparticles per bacterial receptor could be achieved using phages due to the comparatively large surface area of phage, which may accommodate multiple nanoparticles; this property could be useful if bacterial receptors are in low abundance. A related benefit is that the aggregation of nanoparticles with phages on bacteria produces a visible shift in the LSPR spectrum (31), and one might therefore envision applications that combine treatment and detection of bacteria. Second, in addition to the targeting mechanisms evolved by whole phages as described above, chimeric phages can be rationally designed to achieve specificity against different bacterial hosts (8). This potential is largely untapped, as there exists a mostly uncharacterized biological reservoir of phages that could presumably target many different bacterial strains (32). While phages are well known for their host specificity, a number are broad in host range (33), suggesting that the degree of specificity could be tuned depending on the desired application. Third, in practical terms, phages are inexpensive to produce and have typically evolved some hardiness to nonideal environmental conditions. These features make phage-based nanotechnology attractive for biotechnological and biomedical applications.

In this work, we investigated the ability of phage–AuNR bioconjugates (phage–AuNRs; Fig. 1A) to specifically attach to and then kill targeted bacterial cells via the photothermal effect. In this scheme, the phage confers specific targeting while the AuNRs achieve the desired effect. Using chimeric phages we previously engineered (31) to target several bacterial pathogens (two strains of Escherichia coli, Pseudomonas aeruginosa, Vibrio cholerae, and two strains of the plant pathogen Xanthomonas campestris), we first show that the phage–AuNRs can be used to detect specific bacteria through phage-mediated aggregation of AuNRs on the bacterial surface, which causes a red shift of the longitudinal LSPR peak. Next, we use NIR irradiation to induce death of the targeted bacteria via photothermal heating, both in solution and in a P. aeruginosa biofilm grown on a substrate of mammalian epithelial cells. Photothermal lysis was highly selective and resulted in extensive killing of targeted bacteria within minutes, with low rates of damage to nontarget bacteria and mammalian cells (Fig. 1B). After photothermal lysis, the phages are no longer capable of replication, allowing control over dosage in principle. The potential of phage–AuNRs for treatment and diagnosis of antibiotic-resistant bacterial infections is discussed.

Fig. 1. Schematic of phage–AuNR bioconjugates for bacterial detection and cell killing. (A) Filamentous phage M13 (gray) was engineered to express the receptor-binding protein from a foreign phage (blue) fused to wild-type g3p to obtain retargeted chimeric phage. Chemical modification by N-succinimidyl-S-acetylthiopropionate (SATP) introduced thiol groups (yellow) along the phage coat, and gold nanorods (AuNRs) were conjugated to the phage via thiol–gold bonds. (B) Phage–AuNR bioconjugates recognize specific bacteria (blue) in the context of mammalian cells and other nontarget bacteria (green). Attachment of phage triggers AuNR aggregation on the cell surface, resulting in a red-shifted LSPR spectrum (indicated as magenta nanorods) for bacterial detection. Exposure to NIR light induces photothermal heating of AuNRs, leading to highly elevated temperatures localized by the phage, resulting in death of the targeted bacteria.

Discussion We present an antibacterial strategy using phages conjugated to AuNRs (phage–AuNRs, referred to in the following discussion as “phanorods,” a portmanteau of “phage” and “nanorods”). The phages attach to targeted bacteria, and irradiation of the nanorods by NIR light causes LSPR excitation. This energy is released as heat, destroying the phage as well as bacteria bound to the phage. The phanorod strategy has important advantages over traditional approaches to phage therapy. First, phage therapy suffers from the major difficulty of managing a replicating and evolvable entity. While the evolutionary capacity of phages is advantageous for overcoming bacterial resistance against a phage, evolutionary potential is an important biocontainment concern in practice. Second, nonlinear replication dynamics mean that dosages cannot be easily controlled, which may be problematic if cell lysis releases endotoxins triggering deleterious host responses (e.g., septic shock). Phanorods are destroyed during irradiation, preventing replication and evolution during treatment and enabling control over dosage. Irradiation could also be used to inactivate excess phanorods after use, avoiding negative impacts, such as evolution of resistant organisms, currently associated with antibiotics in the waste stream. Third, evolution of resistance is an important challenge for any antibacterial strategy, including phanorods. However, because the phage is used only for attachment to cells and downstream events (e.g., replication) are not relevant, bacterial mechanisms for resistance should be limited to alterations of the receptor, presenting a smaller mutational target for evolution of resistance. Fourth, phanorods serve simultaneously as diagnosis and cytotoxic reagents, as the change in the LSPR spectrum can be used to recognize bacterial species. Therefore, although there may be situations in which therapy with phages per se is desired (e.g., if exponential replication dynamics are needed), phanorod pharmacokinetics and pharmacodynamics may more closely resemble those of a typical drug rather than a living organism, which would be advantageous for most therapeutic situations. Conversely, one may consider how phanorods compare to antibody-conjugated nanorods. In addition to unique search mechanisms (see Introduction), phages can possess very high affinities to their targets. For example, the affinity of M13KE for F+ E. coli is 2 pM (12), which is several orders of magnitude greater than the affinities of antibodies reported against F pilin (47) and also higher than affinities of most antibodies isolated from mice (30 to 500 pM, with those at low picomolar affinity being quite rare) (48, 49). In the experimental comparison done here between M13KE–AuNRs and αLPS–AuNRs, we found that M13KE–AuNRs indeed outperformed αLPS–AuNRs in photothermal ablation experiments, despite the greater amount of LPS present on the cell surface compared to the F pilus. It may be of particular interest to eradicate targeted bacteria in small numbers, such as “persister” cell populations that escape other treatment modalities (50). In such applications, the very high affinities and often high specificities of phages would avoid competition for binding by nontarget cells and thus prevent loss of phanorod activity compared to αLPS–AuNRs, as demonstrated here in a mixed bacterial population. The experiments also confirmed the environmental hardiness of the phanorods, which, unlike the αLPS–AuNRs, tolerated the range of pH tested (pH 3 to 10) as well as heat treatment. We speculate that very high affinities and “hunting” strategies for bacteria, environmental hardiness, and tolerance to mutations and thus chemical modification, which characterize phages, are traits under selection by the ongoing evolutionary “arms race” between bacteria and phages (51). This constellation of traits is advantageous for therapeutic nanomaterials and may be unique to phages. The phanorod strategy presented here is best suited for treatment of directly accessible tissues or surfaces. Near-term applications could be localized topical therapy, particularly for wound infections or colonization of medical devices, in which the phage–nanorods can be directly applied to the biofilm. For example, while P. aeruginosa is well known as a pulmonary pathogen, drug-resistant P. aeruginosa is also a major pathogen in chronic wounds (52), surgical site infections, and burns (53). Such wounds would be directly accessible for application of therapeutic nanomaterials and NIR irradiation. Bacterial biofilms represent a difficult challenge (54) for treatment, as the protective extracellular matrix often inhibits access by antibiotics. However, heat can be transferred without molecular penetration into the biofilm. While the phanorod strategy should be generalizable to different bacterial strains depending on the creation of chimeric phages, we focused on P. aeruginosa as one of three “critical priority” bacterial pathogens identified by the World Health Organization (55). In this work, we demonstrated that phanorods were effective in killing a P. aeruginosa biofilm grown on epithelial cell culture (see SI Appendix, Text S2 for further discussion). Some photothermal damage was incurred by epithelial cells, although the viability measured here is likely a lower bound since the biofilm was in direct contact with the monolayer; underlying cellular layers in a physiological context would likely sustain less damage. The phanorods used here absorb in the relatively biologically transparent window of NIR light. In principle, irradiation could be directed only toward areas where activation of the phanorods is desired, reducing potential side effects. Whether iterative phanorod application could be effective in treating thicker and deeper biofilms (e.g., abscesses) without substantial harm to surrounding tissue is an important practical issue. In addition, while excess phanorods could be removed by washing for certain wounds, a consideration for other applications is the in vivo biodistribution of phanorods in the absence of target bacteria. Nevertheless, phanorods and other alternative strategies merit consideration given the current need to develop new antibiotic agents.

Methods Materials. Reagents were obtained from the following sources: gold(III) chloride trihydrate (HAuCl 3 •3H 2 O) (99.9%; Sigma), sodium borohydride (NaBH 4 ) (98%; Fisher Scientific), trisodium citrate dihydrate (99.9%; Sigma), Escherichia coli (Migula) Castellani and Chalmers (ATCC27065, ATCC), E. coli ER2738 (NEB), Xanthomonas campestris pv. campestris (ATCC33913), Xanthomonas campestris pv. vesicatoria (ATCC35937), Pseudomonas aeruginosa (Schroeter) Migula (ATCC 25102), Vibrio cholerae 0395 (donation from Dr. Michael J. Mahan, University of California, Santa Barbara, CA), M13KE phage (NEB), M13-NotI-Kan construct (12), sodium chloride (NaCl) (99%; Fisher BioReagents), tryptone (99%; Fisher BioReagents), yeast extract (99%; Fisher BioReagents), E. coli ER2738 (NEB), fluorescein-5-maleimide (97%; TCI), N-succinimidyl-S-acetylthiopropionate (SATP) (Thermo Fisher Scientific), hydroxylamine hydrochloride (99%; Sigma), N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC) (99.9%; Sigma), N-hydroxysuccinimide (NHS) (98%; Sigma), mouse monoclonal anti-E. coli LPS antibody ([2D7/1] ab35654; Abcam), 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF) (Invitrogen), 5-bromosalicylic acid (5-BAA) (>98.0%; TCI), SYTO 9, PI (Thermo Fisher Scientific), BODIPY C10 (donation from Dr. M. Kuimova from Imperial College London, London, UK), poly(ethylene glycol) (PEG-8000; Sigma), dialysis kit (MWCO 3500 Da; Spectrum Labs), tetracycline (98%; Fisher Scientific), kanamycin sulfate (98%; Fisher Scientific), ampicillin (98%; Fisher Scientific), isopropyl β-d-1-thiogalactopyranoside (IPTG) (99%; Fisher Scientific), Mix & Go competent cells (Zymo Research), QIAprep Spin Miniprep Kit (Qiagen), QIAquick Gel Extraction Kit (Qiagen), KpnI-HF/NotI-HF restriction enzyme and T4 DNA ligase (NEB), and thiol-PEG-acid (HOOC-PEG-SH; PEG average M n 5,000; Sigma). Chimeric Phages. The construction of the chimeric phages used here [M13-g3p(CTXϕ), M13-g3p(Pf1), M13-g3p(ϕLf), M13-g3p(ϕXv), and M13-g3p(If1)] was previously reported (31). Phages were propagated according to standard protocols. Phage concentrations were quantified by real-time PCR as previously described (31). See SI Appendix for more details. Thiol Functionalization of Phages. Phages were treated with SATP for chemical modification of accessible amine groups of the capsid based on a known protocol (56). See SI Appendix for more details. Synthesis of AuNRs. The AuNRs were synthesized through a modified method by Murray and coworkers (34). See SI Appendix for more details. Preparation of Phage–AuNR Bioconjugates. Conjugation of thiolated phage with AuNRs was conducted based on a known bioconjugation method (39). The AuNRs were resuspended in Tris buffer (50 mM, pH 3). Two hundred microliters of thiolated phage (1 × 1011 pfu/mL) was added dropwise to 1-mL AuNRs solution (6.8 nM). The suspension was incubated at room temperature for 2 h under moderate rotation. The phage–AuNR conjugates were purified by repeated centrifugation/resuspension cycles (8,000 rpm for 30 min, resuspension in 1 mL of water). The reaction was monitored by UV-vis spectrophotometer (Shimadzu UV-1800) and Zetasizer APS (Malvern) to follow changes of LSPR and zeta potential. The trace amount of residual CTAB was further exchanged with HOOC-PEG-SH (2.0 μM) under the same conditions for 24 h. The phage–AuNR bioconjugates were purified by centrifugation and resuspended in 200 μL of water. Visualization of Binding of M13KE–AuNR to E. coli. 1011 M13KE–AuNR bioconjugates (i.e., containing 1011 M13KE phages) were incubated with 2 mg/mL fluorescein-5-maleimide in 1 mL of PBS buffer (pH 7.0) with gentle stirring at room temperature overnight. The free dye was removed by extensive dialysis (MWCO 3,500 Da) in 500 mL of PBS buffer (pH 7.0), and bioconjugates were concentrated by ultrafiltration to ∼200 µL with an Amicon Ultra-4 10000 filter. The M13KE–AuNR bioconjugates were incubated with 1 mL of Top 10F′ cells expressing cyan fluorescent protein at an optical density ∼0.6 for 30 min at room temperature (12). Free bioconjugates were removed by centrifugation at 5,000 rpm and discarding the supernatant. The pellet was washed by PBS buffer twice and resuspended in 1 mL of PBS buffer for microscopy. The fluorescence images were recorded on a Leica SP8 confocal microscope (Leica), with excitation at 405 nm. Detection of Bacteria with Phage–AuNR. Bacterial cells (strains of E. coli, V. cholerae, P. aeruginosa, and X. campestris listed above) were grown in liquid culture and cell concentrations estimated by optical density as previously described (31) (SI Appendix, Methods). The cells were collected by centrifugation and resuspended in MilliQ water at the desired concentrations. Fifty microliters of bacterial solution was added into 100 µL of phage–AuNR bioconjugate solution (∼1011 phage particles per mL) in 1.5-mL tubes. The absorbance of the solutions was recorded by UV-vis spectroscopy (Shimadzu UV-1800) after a 30-min incubation at room temperature. The specificity of detection was tested by adding a different bacterial species or strain from the host of the phage source of the g3p-N homolog (Table 1), or by adding a mixture of host cells [E. coli (F+), V. cholerae 0395, E. coli (I+), X. campestris (pv campestris), X. campestris (pv vesicatoria), and P. aeruginosa] for the assay. Photothermal Lysis of Bacteria with Phage–AuNRs. To test thermolysis in aqueous solution, the mixture of bacterial solution and phage–AuNRs (1011 phages, and 106 cells in 1 mL solution) was irradiated for various time periods (0 to 10 min) using an 808-nm diode laser (3.0 W/cm2; Q-BAIHE Company) ∼8 cm from the top of the solution. After irradiation, a 10-μL aliquot of the resulting solution was diluted into PBS buffer (0.1 M, pH 7.4) to a ratio of 1:1,000 and cultured on plates for colony counting using ImageJ. X. campestris (pv campestris) and X. campestris (pv vesicatoria) were cultured on YPD agar plates with no antibiotics; the other cells were cultured on LB plates. E. coli ER2738 was cultured on LB plates containing 10 µg/mL tetracycline. The E. coli strains expressing cyan or citrine fluorescent proteins (12) were cultured on LB plates containing 1 mM IPTG and 100 µg/mL ampicillin. To test thermolysis of a P. aeruginosa biofilm grown on solid support, the biofilm was prepared using a protocol modified from the literature (41). A single colony of P. aeruginosa was selected and grown in LB overnight at 37 °C in a shaker incubator. The overnight culture was diluted 100-fold into fresh medium. One hundred fifty microliters of the dilution was added to Lab-Tek plates (culture area, 0.7 cm2) and incubated overnight at 37 °C. After incubation, the liquid was removed by turning the plate over and shaking out the liquid. The remaining biofilm was washed by submerging the plate in a small tub of water and shaking out water twice. Three hundred microliters of M13-g3p(Pf1)–AuNR bioconjugates (1013 phages per mL) were added into the biofilm and incubated for 30 min. Unbound bioconjugates in suspension were removed by pipetting. The biofilm was irradiated with the NIR laser for 10 min as described above. Cell viability was studied by growing the resuspended bacteria on LB plates (1 µL of cell suspension was diluted in 1 mL of PBS buffer, and 5 µL of the dilution was plated onto LB plates) for colony counting, and by confocal microscopy with live/dead cell viability staining with SYTO9 and PI. To test thermolysis of a P. aeruginosa biofilm grown on mammalian epithelial cells, wild-type MDCKII epithelial cells in suspension were seeded at 5.0 × 104 cells per well in eight-well chamber slide Lab-Tek plates and grown to confluency. Dulbecco’s modified Eagle’s medium (DMEM) (Thermo Fisher; 11885076) was supplemented with 10% FBS (Thermo Fisher; 10437036) and 1% penicillin–streptomycin (P/S) (Thermo Fisher; 15140122), and kept in an incubator at 37 °C with 5% CO 2 . After 48 h, the culture medium was removed, and the cells were washed twice with PBS buffer to remove traces of P/S. The biofilm of P. aeruginosa was cultured on the top of the MDCKII epithelial cells in 15 mM Hepes buffer (pH 7 with DMEM and 10% FBS) as described above. Planktonic cells were removed from the biofilm by careful pipetting. Incubation with M13-g3p(Pf1)–AuNR (1013 phages per mL) and NIR irradiation of the bacterial cells was performed as described above. Cell viability of the MDCKII cells was assayed by the PrestoBlue cell viability assay, and bacterial and mammalian cells were assessed by confocal microscopy with live/dead cell viability staining with SYTO9 and PI. Cytotoxicity of Phage and Phage–AuNR. The PrestoBlue cell viability assay was performed according to the manufacturer’s protocol (Invitrogen). MDCKII epithelial cells were cultured to confluency in a 96-well microtiter plate as described above. After 24 h, the medium was replaced by 180 µL of fresh DMEM (containing 10% FBS) with 20 µL of samples of various concentrations of M13KE phage or phage–AuNR bioconjugates in PBS buffer. The cells were incubated for another 48 h, and the medium was replaced by 90 µL of fresh medium, mixed with 10 µL of PrestoBlue. The cells were incubated at 37 °C for 30 min, and fluorescence signals were measured on a TECAN infinite 200 Pro plate reader (TECAN). The excitation wavelength was 560 nm (bandwidth, 9 nm), and the emission wavelength was 600 nm (bandwidth, 20 nm). Similar results were obtained using a 1-h incubation time. The cell viability was expressed as a percentage relative to the control cells (MDCKII cells incubated with PBS buffer with no phages or phage–AuNR under the same conditions). Temperature Measurement of Cells Using BCECF. To measure the temperature of suspended cells during thermolysis, E. coli ER2738 cells attached to the M13KE–AuNR bioconjugates (prepared as described above) were incubated with 10 µM BCECF solution at room temperature for 20 min. The free dye was removed by centrifugation. The cells were washed three times and resuspended in PBS buffer (0.1 M, pH 7.4). The bulk fluorescence of BCECF (λ ex = 490 nm, λ em = 500 to 600 nm) in the sample at room temperature was determined using a Fluoromax-4 spectrofluorometer (Horiba). The presence of AuNRs did not affect the observed spectrum of BCECF (SI Appendix, Fig. S9E). A standard solution of BCECF (0.15 µM in PBS buffer) was prepared to match the observed bulk fluorescence at room temperature. The fluorescence spectrum of the standard BCECF solution was measured at varying temperatures using a Peltier-based cuvette holder (Horiba) to construct a standard curve. The fluorescence of the E. coli–phage–AuNR suspension was recorded simultaneously with laser irradiation from above the sample to induce thermolysis. To estimate the temperature of cells in a biofilm during thermolysis, a calibration curve was first obtained by recording fluorescence images of a biofilm stained with BCECF at different temperatures (from 25 to 90 °C) using a Lauda Eco RE415 Silver cooling thermostat system (LAUDA-Brinkmann) with a confocal microscope (excitation at 488 nm; TCS SP8; Leica Microsystems). The fluorescence intensities were calculated using ImageJ to obtain the calibration curve between intensity vs. temperature. Cells were identified from background by thresholding and intensities recorded from the “Analyze Particles” tool in ImageJ. Threshold settings were kept identical between images. Between measurements, the microscope laser was shut down, and the sample was covered with aluminum foil to reduce photobleaching while the temperature was adjusted. A P. aeruginosa biofilm was incubated with M13-g3p(Pf1)–AuNR bioconjugates and stained by BCECF as described above. The free dye was removed by pipetting off the liquid phase and washing three times with PBS buffer. The biofilm was irradiated by NIR light for 10 min as described above, and a fluorescence image was taken immediately. Viscosity of Cell Membranes Measured by Molecular Rotor BODIPY C10. The viscosity of mammalian and bacterial cell membranes during thermolysis was measured by staining the biofilm with BODIPY C10 (incubation with 10 µM BODIPY C10 at 37 °C for 20 min followed by replacement of DMEM) and measuring by FLIM before and after NIR laser irradiation (described above). Fluorescence lifetime images were recorded using a time-correlated single-photon counting card (Leica Falcon FLIM). Imaging was achieved using a confocal microscope (TCS SP8; Leica Microsystems) with a Leica SuperK white light laser, which provided pulsed excitation permitting time-resolved fluorescence imaging. The excitation wavelength used was 488 nm. The data were analyzed by LAS X FLIM/FCS software (Leica Falcon FLIM). The bacteria were identified manually by morphology of the cells in confocal microscope images. The viscosities were calculated from FLIM data according to the following viscosity-lifetime calibration equation, which was obtained by measuring the fluorescence lifetime of BODIPY C10 in different solutions of methanol/glycerol mixtures with known viscosities (43): log ⁡ V = log ⁡ T + 0.75614 0.4569 . [1] Here, V is viscosity (in centipoises), and T is fluorescence lifetime (in nanoseconds). Concentration of AuNRs. Single-particle ICP-MS was performed with an Agilent 7900 ICP-MS (Santa Clara) to determine the concentration of the AuNRs. The phage–AuNR bioconjugates were incubated with 5% nitric acid for a week to degrade the virus before the measurement. The analysis was carried out in a time-resolved analysis mode with an integration time of 100 µs per point and no settling time between measurements. The data analysis was conducted with the Agilent ICP-MS MassHunter software (version C.01.04 Build 544.3) via single-nanoparticle application module. TEM. TEM was performed on a Tecnai FEI G2 Sphera microscope (Materials Research Laboratory [MRL], University of California, Santa Barbara) as previously described (31). M13KE–AuNR samples were prepared as described above (Methods, Visualization of Binding of M13KE–AuNR to E. coli). The AuNR size was calculated by measuring 200 AuNRs. Zeta Potential Measurements. Zeta potentials were measured by using a Malvern Zetasizer Nano ZSP operating a 4 mW He–Ne laser at 633 nm as previously described (31). Data from three or more individual samples were averaged, and each sample was measured five times (10 runs each). See SI Appendix, Methods for description of additional optical characterization, synthesis of AuNRs, colocalization analysis, antibody–AuNR experiments, chimeric phages, phage and bacterial propagation, and EPS quantitation. Data Availability. All data are included in the manuscript and SI Appendix.

Acknowledgments Financial support from the NIH (DP2 GM123457-01 to I.A.C.), the Institute for Collaborative Biotechnologies (Contract W911NF-09-0001 from the US Army Research Office), and the National Science Foundation (CMMI 1662431 to B.L.P.) is acknowledged. We thank M. Mahan for bacterial strains and advice, W. J. Nelson for the MDCKII cell line, and Petra Levin for advice on bacterial biofilms. We thank M. Kuimova for kind provision of molecular rotor BODIPY-C10. We acknowledge the use of the Neuroscience Research Institute–Department of Molecular, Cellular, and Developmental Biology Microscopy Facility and the resonant scanning confocal microscope supported by the NSF Major Research Instrumentation Grant DBI-1625770; the Biological Nanostructures Laboratory (UV-vis) within the California NanoSystems Institute, supported by the University of California, Santa Barbara, and the University of California, Office of the President; ICP-MS instrumentation from the Keller Laboratory at University of California, Santa Barbara; and the MRL Shared Experimental Facilities (attenuated total reflection–FTIR, TEM, XPS) supported by the Materials Research Science and Engineering Center Program of the NSF under Award DMR 1720256, a member of the NSF-funded Materials Research Facilities Network (https://www.mrfn.org/).

Footnotes Author contributions: H.P. and I.A.C. designed research; H.P. and R.E.B. performed research; H.P., L.P.D., and B.L.P. contributed new reagents/analytic tools; H.P. and I.A.C. analyzed data; and H.P. and I.A.C. wrote the paper.

Competing interest statement: A provisional patent application has been filed: University of California, Santa Barbara case 2018-758.

This article is a PNAS Direct Submission.

This article contains supporting information online at https://www.pnas.org/lookup/suppl/doi:10.1073/pnas.1913234117/-/DCSupplemental.