Engineered T4 Viral Nanoparticles for Cellular Imaging and Flow

ARTICLE pubs.acs.org/bc

Engineered T4 Viral Nanoparticles for Cellular Imaging and Flow Cytometry Kelly L. Robertson, Carissa M. Soto, Marie J. Archer, Onyekachi Odoemene, and Jinny L. Liu* Contributed from the Center of for Bio/Molecular Science and Engineering, Naval Research Laboratory, 4555 Overlook Avenue SW, Washington, DC 20375, United States

bS Supporting Information ABSTRACT: Viruses are of particular interest as scaffolds for biotechnology applications given their wide range of shapes and sizes and the possibility to modify them with a variety of functional moieties to produce useful virus-based nanoparticles (VNPs). In order to develop functional VNPs for cell imaging and flow cytometry applications, we used the head of the T4 bacteriophage as a scaffold for bioconjugation of fluorescent dyes. Bacteriophage T4 is a double-stranded DNA virus with an elongated icosahedron head and a contractile tail. The head is ∼100 nm in length and ∼90 nm in width. The large surface area of the T4 head is an important advantage for the development of functional materials since it can accommodate significantly larger numbers of functional groups, such as fluorescent dyes, in comparison with other VNPs. In this study, Cy3 and Alexa Fluor 546 were chemically incorporated into tail-less T4 heads (T4 nanoparticles) for the first time, and the fluorescent properties of the dyeconjugated nanoparticles were characterized. The T4 nanoparticles were labeled with up to 19 000 dyes, and in particular, the use of Cy3 led to fluorescent enhancements of up to 90% compared to free Cy3. We also demonstrate that the dye-conjugated T4 nanoparticles are structurally stable and that they can be used as molecular probes for cell imaging and flow cytometry applications.

’ INTRODUCTION Recently, the use of viral nanoparticles (VNPs) as materials for biotechnology applications has gained interest given their biocompatibility, the diversity of shapes and sizes, and the possibility to tailor their surface characteristics.1-4 VNPs produced from plant and animal viruses and bacteriophages have been used in biosensing, imaging, vaccine development, and drug and gene delivery.3-7 Of these, bacteriophages (or phages) are among the most promising candidates as scaffolds in biotechnology applications due to their robust nature and unique display system, which enables the expression of a wide variety of peptides and polypeptides rendering diverse surface functionality.3,8,9 The phage head varies in shape and size from micrometer-size in the filamentous type (fd, M13) to a few hundred nanometers in the spherical (PhiX174, G4, S13) and icosahedral type (T7, T4, MS2) and the tail, when present, can be contractile or flexible.3,10,11 Bacteriophage T4 has several characteristics that distinguish it from other phage species and make it an ideal scaffold for biotechnology applications. For instance, it has a flexible and unique display system12-14 that has been demonstrated to display peptides and full-length proteins on the viral capsids. Combining this feature with the possibility to engineer its morphology15 significantly increases the range of functional materials that can be devised. In addition, when functionalized through chemical conjugation, the large surface area of the head allows a higher loading capacity and spatial distribution of the functional groups, which presumably leads to higher sensitivity.16 r 2011 American Chemical Society

The size and shape are also advantageous, because it has been demonstrated that cellular uptake occurs preferentially with spherical particles with sizes between 70 and 100 nm over elongated ones even with the same dimensions.17 Figure 1a presents a schematic representation of bacteriophage T4 consisting of an icosahedral head, a contractile tail, and tail fibers, while Figure 1b shows the concept of a T4 nanoparticle obtained through genetic engineering. Deletion of the tail and its fibers results in nanoparticles with dimensions of ∼100 ∼90 nm2, 18,19 which can be functionalized with fluorescent dyes using standard bioconjugation techniques. Bacteriophage T4 has been proven useful in sensor probes20-22 and vaccine carriers,23-25 and the use of tail-less T4 nanoparticles (T4 NPs) as detection elements has been suggested,26 yet no practical applications have been reported to date. We previously used atomic force microscopy to examine the variables that influence tail-less T4 NP assembly on surfaces.19 In the work presented here, we demonstrate, for the first time, the use of T4 NPs as fluorescent probes for cellular imaging and flow cytometry applications. The properties of the T4 NP give it the potential of becoming a viable scaffold for biomaterial applications, but we are currently limited by the scarcity of information on its surface chemistry. The large size of the T4 head makes it Received: August 10, 2010 Revised: January 4, 2011 Published: March 04, 2011 595

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dye-T4 NPs can be used to visualize cells and remain inside cells for at least 72 h.

’ EXPERIMENTAL SECTION Purification of Tail-Less T4 Head Scaffolds. T4 K10 (38-

-

51 denA- denB-), a kind gift from Dr. Lindsay Black at University of Maryland at Baltimore Medical School, was first propagated in the suppressor Escherichia coli host strain, CR63, to obtain the infectious phages. The infectious phages were then used to produce noninfectious tail-less T4 nanoparticles (T4 NPs) using the nonsuppressor host E. coli strain, Rosetta, modified from a previous procedure.40 In brief, Rosetta grown in M9S (OD600 = 0.5) supplemented with 1/3 volume of Luria Broth were infected with K10 phage at a multiplicity of infection (MOI) = 3, followed by a repeated infection after 9 min and continued incubation for 2 h at 37 °C. Cells were spun down at 10 000 g for 10 min and resuspended in 50 mM potassium phosphate (pH 7.5) supplemented with 10 mM MgCl2 and 2 mM CaCl2; CHCl3 (1/20 of total volume), DNase I (40 μg/mL), RNase I (50 μg/mL), and 0.4 μg of PMSF (phenylmethanesulfonylfluoride or phenylmethylsulfonyl fluoride) were then added to the cell suspension and the cells were gently shaken at 37 °C for 1-2 h. The cell debris was removed by spinning at 15 000 g for 30 min. The cell lysate containing the head scaffolds was concentrated through a Microcon YM-100 membrane according to the manufacturer’s procedure (Millipore Corp, MA). The membrane was then washed 3-6 times with 50 mM potassium phosphate buffer, pH 7.5, supplemented with 10 mM MgCl2 (50 KP/ 10 MgCl2) and 0.1% sodium azide. The T4 NPs were then purified by gel filtration with Superose 6 (GE Healthcare Biosciences, NJ). The flowthrough and eluted fractions were then assessed using a 1.2% Tris-acetate agarose gel stained with either ethidium bromide or Coomassie blue. Dye Conjugation. Tail-less T4 NP solutions in 50 KP/10 MgCl2 and 0.1% sodium azide were desalted using a pre-equilibrated HiTrap Desalting Column (GE Healthcare Biosciences, Piscataway, NJ) and eluted fractions were analyzed by UV-vis spectroscopy (Cary 5000, Varian). The second 1.5 mL elution contained most of the T4 NPs. For the reaction optimization experiments, T4 NPs were reacted with fluorescein-5-isothiocyanate (FITC; Invitrogen, Carlsbad, CA) and Alexa Fluor 488 sulfodichlorophenol ester (Alexa488-SDP ester, Invitrogen) in various buffer conditions. When developing an optimally fluorescent dye-T4 NP, 9.0 1010 K10 heads (9 μg of protein) in 10% DMSO (Sigma-Aldrich, St. Louis, MO) were reacted with Cy 3 Mono NHS ester (Cy3-NHS; GE Healthcare Biosciences) or Alexa Fluor 546NHS ester (Alexa 546-NHS; Invitrogen, Carlsbad, CA). Molar ratios of dye to T4 NP lysines were 0.1 (for Alexa Fluor 546 only), 0.6, 1.1, 1.5, 2.0, 2.8, 5, 10, 13, and 20. Prior to the addition of DMSO and dye, T4 NPs in 50 KP/10 MgCl2 were incubated on ice for 5 min after which DMSO was added slowly with gentle mixing. Then, corresponding amounts of 10 μg/μL NHS-Cy3 or NHS-Alexa Fluor 546 in DMSO were added, and the mixture was vortexed. T4 NPs were kept at a concentration of 5.8 1011 particles/mL in the reaction mix, and the mixture was incubated at room temperature in the dark for 16 h. From these reactions, we obtained a series of samples containing a different number of dyes per T4 NP. Dye-labeled T4 NPs were loaded separately into a Superose 6 prep grade (GE Healthcare Biosciences) column pre-equilibrated in 50 KP/ 10 MgCl2/0.1% sodium azide. Individual 1 mL elutions were collected and analyzed by gel electrophoresis, atomic force microscopy, and UVvis spectroscopy. For gel electrophoresis, 10 μL of unmodified and dyelabeled T4 NPs were loaded into a 1.2% agarose gel. The gel was visualized without staining using HP Scanjet 5590 scanner. DNA and protein visualization was also performed on the same gel using ethidium bromide and Coomassie blue staining, respectively. Images from

Figure 1. Experimental strategy for probe synthesis. (a) Schematic representation of bacteriophage T4. (b) Schematic representation of a tail-less T4 NP obtained by deletion of the tail using genetic engineering; functional T4 NPs can be developed by incorporating fluorescent dyes using standard bioconjugation techniques.

difficult to grow single crystals for X-ray crystallography at a resolution sufficient for determining the exact location of amino acids on the surface. The most current structure available comes from cryo-electron microscopy at a 2.2 nm resolution.27 Thus, to further demonstrate its potential as an addressable nanobuilding block and understand its surface chemistry, the reactivity of the surface amines was investigated. While lysine groups are typically used to couple biomolecules due to their abundance and wellknown chemistry, other amines may be available for conjugation as well. The total number of lysines on the T4 head is 32 303 (Supporting Information Table S1). In the work presented here, we selected Cy3 and Alexa Fluor 546 as model dyes and characterized their performance on the T4 NP. These two fluorophores, while sharing similar excitation/ emission spectra, have been reported to exhibit strikingly different performance characteristics, which are thought to be due, in part, to their molecular properties, but also to their neighboring interactions.28,29 It is therefore relevant to understand their behavior on the T4 scaffold under a particular set of experimental conditions. Chemical bioconjugation of organic dyes has been utilized to prepare fluorescent probes using plant viruses such as cowpea mosaic virus (CPMV),16,30-33 turnip yellow mosaic virus (TYMV),34 tobacco mosaic virus (TMV),35,36 and potato virus X (PVX);37 however, there are fewer examples of the use of bacteriophages as scaffolds for this purpose.38 In plant viruses containing small capsids, such as CPMV and TMV (30 and 28 nm in diameter, respectively), the number of dyes that can be incorporated is limited by the surface area of the capsids and corresponding reactivities of solvent-accessible amino acids on the virus surface. Precise control of the spacing between the dyes becomes more critical within this limited surface area in order to avoid dimer formation and quenching of the dyes.32 While TMV with its “hollow-rod” shape has a larger surface area, the chemical conjugation is cumbersome, since commonly used residues, such as lysines, are not exposed on the surface.35,39 PVX on the other hand has an elongated capsid (∼500 nm18 nm), and even with this significantly larger surface area, the maximum number of dyes incorporated is only 1600 per capsid through primary amine modification.37 Here, we demonstrate that more than 1.9 104 dyes can be incorporated on a single T4 NP; to our knowledge, this is the highest number of dyes incorporated onto a single VNP. Our results also show that the T4 probes are so efficient that even those with dye numbers as low as 350 dyes per particle give good fluorescence intensities for in vitro applications. In addition, the 596

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Bioconjugate Chemistry ethidium bromide staining were obtained using UVP Bio Doc-It system, a UV transilluminator. Coomassie blue stained images were captured by the HP scanner. Spectroscopic Measurements. UV-vis spectroscopy was performed to determine the amount of T4 NPs and dye present in the reaction samples. Protein concentration was determined using a Bradford assay according to the manufacturer’s procedure (Bio-Rad, Hercules, CA), and the number of T4 NPs per sample was then determined using the relationship that 1 mg of purified NPs is equivalent to 9.70 1012 tail-less T4 heads. The amount of Cy3 and Alexa 546 in the T4 solution was determined from absorbance at 550 and 555 nm, respectively, and by a standard curve produced using each free dye. UV-vis measurements were performed using Varian Cary-5000 UV-vis-near-IR spectrometer with Cary Win UV Scan Application v 3.00 software. Fluorescence spectroscopy was also carried out to determine the fluorescence intensity (F.I.) of dye-T4 NPs. The Cy3-T4 and Alexa 546-T4 NPs were excited at 550 nm. The measurements were taken on a Cary Eclipse fluorescence spectrophotometer. To calculate the percentage of fluorescent enhancement, the following equation was used: ([dye-T4 complex F.I.] - [F.I. of free dye])/[dye-T4 complex F.I.] 100. Atomic Force Microscopy (AFM). For AFM characterization, 5 μL of the unmodified or dye-labeled T4 NPs were sonicated for 510 s, deposited on freshly cleaved mica, and kept in a humid environment at room temperature for 12 h. After adsorption of the T4 NPs onto the mica surface, the substrates were allowed to dry at room temperature, carefully rinsed with Milli-Q water, and allowed to dry at room temperature once more. Unmodified and conjugated T4 NPs were characterized using a Multimode scanning probe microscope (Veeco Instruments, Plainview, NY, USA) operated under tapping mode in air with a silicon cantilever (Nanoscience Instruments, Phoenix, AZ), 125 mm long, with an apex curvature radius of 5-6 nm, a resonant frequency of 300 kHz, and a spring constant of 40 N/m. The scanning rate was 0.5 Hz, at 0° angle. Image processing was performed using Research NanoScope II software ve 7.20. All images were filtered using the flattening built-in tool from NanoScope II software. Length and height values of the nanoparticles were obtained by utilizing the built-in tool for cross section analysis. Cell Culture. A549 (American Type Culture Collection) lung cancer epithelial cells were grown at 37 °C under 5% CO2 atmosphere in Dulbecco’s modified Eagle’s medium (DMEM), supplemented with 10% fetal bovine serum (FBS) (Cellgro, VA) and 1% (v/v) penicillin/ streptomycin (Sigma, CA), and detached from culture flasks using trypsin-ethylenediamine-tetraacetic acid (EDTA) (Cellgro, VA). After the cells were harvested, they were counted and seeded onto microscope slides, 6-well tissue culture treated plates, or 24-well tissue culture treated plates and allowed to attach overnight. Confocal Microscopy. A549 cells on microscope slides were treated with the dye-T4 NPs at varying times and concentrations in DMEM supplemented with 1% penicillin/streptomycin. The cells were then washed three times for 5 min each with 1 PBS (phosphate buffered saline). The cells were fixed with 4% paraformaldehye in 1 PBS for 15 min and then washed twice with 1 PBS. The cells were then stained with 500 μM 40 ,6-diamidino-2-phenylindole (DAPI) in 1 PBS for 5 min and washed three times with 1 PBS. Confocal microscopy was performed using a Nikon Eclipse TE2000 Confocal Imaging System, λexe = 402 nm (DAPI) and 561 (Cy3/Alexa 546). Images were taken with a 60 objective and are of integrated Z-stacks with 8 steps, 0.8 μm step size, and 10 averages per step. Flow Cytometry. A549 cells on 24-well tissue culture plates were treated with 1 105 dye-T4 NPs per cell and analyzed at 4, 8, and 24 h. Following incubation, the cells were washed twice with 1 PBS and harvested with trypsin-EDTA. The cells were then washed once with 1 PBS and treated with a Live/Dead Fixable Far Red Dead Cell Stain

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Kit (Invitrogen, CA) according to the manufacturer's protocols. Following staining, the cells were washed twice in 1 PBS, resuspended in 1 PBS, and run on an Accuri C6 flow cytometer using standard lasers and filters for PE (FL-2) and APC (FL-4). For each sample, 2 104 events were collected in a gate corresponding to the cell population. To determine the percentage of live cells, a scatter plot showing the intensity of the Live/Dead reagent (FL-4) vs forward scatter (FSC) was gated for the population with the lowest fluorescence (live cells). The percentage of cells positive for the dye-T4 NPs was calculated using the histogram subtraction tool on FSC Express v 3. Cell Tracking. A549 cells on 6-well tissue culture treated plates were treated for 6 h with 1 105 A546-T4 NPs per cell. The cells were then washed two times with 1 PBS and harvested with trypsin-EDTA every 24 h up to 96 h. After harvesting, the cells were washed once with 1 PBS and resuspended in 1 PBS for counting and flow cytometry. The Nexcelom Cellometer Vision (Nexcelom, MA) with an optics module for Alexa 546 (VA-595-501) was used to count and measure the percentage of cells that contained A546-T4 NPs. Each time point was done in triplicate.

’ RESULTS AND DISCUSSION Conjugation of Dyes on T4 NPs. The T4 phage head offers a spherical viral capsid that is similar in geometry to other widely used viral nanoparticles, but with the advantage of having a larger surface area for functionalization with dyes, proteins, antibodies, and inorganic nanoparticles. Large spherical viral capsids offer well-defined geometry, are relatively rigid, and offer better spatial distribution of conjugated functional groups. In order to create a noninfectious nanoparticle (NP) for various biomedical applications, the T4 bacteriophage was expressed without the tail (Figure 1b). The tail-less T4 NPs were purified from E. coli, and 3 mg T4 NPs/L of bacteria was recovered with >90% purity. We then used the purified T4 NPs to explore the accessibility of the amine groups to dye modification. There is no information about the location or accessibility of the individual amino acids on the T4 NPs; therefore, by determining the number of reactive amines one can assess its utility for the applications presented here and as a potential scaffold for further chemical conjugation with antibodies, proteins, and targeting molecules. Reaction conditions were selected based on the stability of the T4 NP and the reactivity of dyes in the selected conditions. In general, reactions carried out in buffer alone resulted in an average of 10 lower reactivity in comparison to reactions carried out in 90:10 buffer to DMSO, due to the enhanced solubility of the dye in DMSO. In addition, including MgCl2 (1, 2, or 10 mM) in the reaction buffer enhanced the stability of T4 and did not affect the reactivity of the dyes. In early optimization reactions, fluorescein-5-isothiocyanate (FITC) and Alexa Fluor 488 sulfodichlorophenol ester (Alexa488SDP ester) were compared to determine the level of reactivity of the T4 NP amines in different buffer conditions. Both dyes have similar spectral characteristics, which facilitates the comparison of UV-visible data when determining number of dyes per T4 NP. FITC is known to provide reasonable specificity toward the ε-amine of lysine,30,41 while Alexa 488SDP ester reacts with all primary amines in a similar manner as common NHS-ester dyes. By comparing FITC with Alexa 488-SDP, one can derive conclusions about the availability of amines on the T4 surface. FITC demonstrated low labeling efficiency when the reaction was performed in the optimal T4 NP buffer (50 mM potassium 597



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not observed, demonstrating the efficient purification of the dyeT4 NPs from free dye. Figure 2b shows a gel stained with ethidium bromide. The major band under UV irradiation indicates that the DNA is inside an intact NP. Figure 2c shows an agarose gel stained with Coomassie blue, which stains the intact T4 NPs. The fact that the bands in lanes 2 and 3 correspond to the same bands in all three visualization methods indicates that indeed the dyes are coupled to intact T4 NPs. When comparing the electrophoretic mobility of the unlabeled-T4 NP control (lane 1) and the dye-T4 NPs (lanes 2-3), it can be observed that the dye-T4 NPs have a net charge of higher negative character than the control, since both complexes ran farther into the agarose gel. This effect has been observed in the past with fluorescent dyes and is most likely due to the addition of negative charges associated with the binding of the dyes.42 Both Cy3 and Alexa Fluor 546 NHS esters have a net negative charge (Supporting Information Figure S1) and eliminate one positive charge upon reacting with each amine group on the T4 surface. In addition to the agarose gel, atomic force microscopy (AFM) topographical images were used to examine the structural features of the unlabeled and dye-labeled T4 NPs. Figure 2d-f shows that the T4-NPs with defined shapes and borders are welldispersed. Cross-sectional analysis found that the average length of the unlabeled nanoparticle is 169 ( 14 nm, which is larger than the predicted length of 119.5 nm from the cryo-EM model.43 Its average height is 39 ( 3 nm, half of what was predicted from cryo-EM (Figure 2d). The difference in length is mainly due to both virus collapsing and AFM tip effect; however, the differences in height of the T4 NPs prepared here can be explained by the loss of DNA from the NP during preparation. According to our solution spectroscopic analysis of the T4 NPs, 10% of the capsids still contain DNA. The empty NPs are collapsed to half of the original height during drying for AFM measurements and therefore are most likely flattened on the surface, which may also extend the length. Slight differences in the different labeled T4 NPs are most likely caused by varying degrees of collapse and flattening. Overall, the height and length measurements of the unlabeled and dye-T4 NPs are comparable. The observed shape and close values of the average length and height between the unmodified and the modified T4 NPs indicate that the capsid withstands the conditions used for the chemical conjugation and purification. The relevance of assessing the structural stability of the T4 NPs after modification finds its rationale in the possible effects that dimethylsulfoxide (DMSO) can have on the cellular membrane and protein denaturation.44,45 Although this solvent is widely used in many biology and molecular biology processes, DMSO can denature protein complexes such as the T4 NP. In the present study, the DMSO concentration was kept at 10%; however, we observed significant degradation of the scaffold at higher concentrations of DMSO (data not shown). This degradation is usually evident under AFM imaging as a layer of debris consisting of small clusters of material of irregular shape and size, as well as the presence of DNA. While other spherical virus capsids are less stable in the absence of DNA or RNA, the T4 NPs are stable under our reaction conditions as shown by gel electrophoresis and AFM.46 Spectroscopic Properties of Dye-T4 NPs. UV-vis measurements were taken of the Cy3-T4 and Alexa-T4 NPs and compared to spectra from the free dyes and unlabeled T4 NPs. Figure 3 shows T4 NP samples labeled with Cy3 and Alexa 546. The 260-280 nm peaks in the T4 spectra represent the

Figure 2. Characterization of the T4 NPs. Left: Agarose gel of T4 NPs visualized under (a) visible light with no staining, (b) UV irradiation after ethidium bromide staining, (c) visible light after Coomassie blue staining. Lane M: DNA marker. Lane 1: Unlabeled-T4 NP control. Lane 2: Alexa 546-T4 NP. Lane 3: Cy3-T4 NP. Right: Atomic force microscopy (AFM) topographical images of (d) unlabeled-T4 NPs, (e) Cy3T4 NPs, (f) Alexa 546-T4 NPs. For AFM, 1.5 109 particles were deposited on freshly cleaved mica.

phosphate pH 7.5, 10 mM MgCl2, 10% DMSO) instead of the manufacturer's recommended buffer (100 mM sodium bicarbonate, pH 9.0, 10% DMSO). However, the Alexa 488-SDP reacted similarly in both the optimal T4 NP buffer and its recommended buffer (100 mM sodium bicarbonate, pH 8.3, 10% DMSO). In order to compare the relative reactivity of FITC and Alexa 488SDP ester, a series of reactions were performed in the recommended buffers for each dye. Our results showed a 30% higher load of Alexa 488 in comparison to FITC (data not shown). This result indicates that the SDP ester is labeling additional amines in comparison to FITC, presumably R-amines, since FITC is known to provide reasonable specificity toward ε-amines of lysines. On the basis of this information, for our current studies we used NHS ester-dyes, which are similar in reactivity to the SDP ester, but are available in a wider variety of dyes with different spectral characteristics. Cy3 NHS ester and Alexa Fluor 546 NHS ester dyes were reacted with T4 NPs at various molar ratios of dye to lysine to produce NPs with a range of dyes/virus (D/V). Characterization of Dye-T4 NPs. To analyze the electrophoretic mobility and structural integrity of the T4 NPs after dye attachment and corresponding purification, the NPs were analyzed using agarose gel electrophoresis and atomic force microscopy (AFM). Figure 2 shows three views of an agarose gel that allows us to characterize different aspects of the Dye-T4 NPs. In Figure 2a, the gel has not been stained and the pink bands correspond to the dye conjugated on the NP. This indicates that the dye is coupled to the NP, since the band is running at the position expected for the unlabeled-T4 NP. Free dye, which is expected to run at a position closer to the bottom of the gel, was 598



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which is known to indicate cyanine dye-protein coupling.47 Alexa 546-T4 NPs also show a slight shift in the dye absorbance compared to free Alexa 546, which is typical of other Alexa fluorophores.48 Dye per Virus (D/V) Measurements. The absorbance measurements taken at the dye wavelengths for each dye-T4 NP sample along with the number of NPs present in each sample were used to calculate the D/V. As the molar excess of dyes to lysines was increased, the D/V increased up to 1.9 104 Alexa 546 dyes at 20 dyes to T4 NP lysines (Supporting Information Figure S2). This D/V ratio is 2 orders of magnitude higher than other iscosahedral viruses, such as CPMV30 and 1 order of magnitude higher than the maximum D/V on the rod-shaped potato virus X and tobacco mosaic virus.35,37 This high degree of labeling is possible due to the large number of amine groups present on the T4 NP (Supporting Information Table S1). Fluorescence Properties of Dye-T4 NPs. Fluorescence emission of the dye-T4 complexes was studied as a function of D/V. As shown in Figure 4a and Supporting Information Table S2, fluorescence emission increases as the number of dyes in the Cy3-T4 is increased up to 715 D/V (orange line Figure 4a); beyond that point. fluorescence decreases as the number of dyes increases presumably due to quenching. In contrast, for the Alexa546-T4 series (Figure 4b) the best fluorescence output is obtained at 2166 D/V (orange line). To further analyze this phenomenon, the individual samples were compared to corresponding free dye in solution. Figure 5 shows the fluorescence enhancement obtained from the dye-T4 complexes calculated relative to a comparable amount of free dye in solution. All of the Cy3-T4 NPs examined showed enhanced fluorescence vs free Cy3 dye (Figure 5a) indicating that conjugation to the T4 changes the fluorescence characteristics of the dye. However, the enhancement of the Cy3-T4 NPs decreases linearly as the number of D/V increases (r = 0.99). In contrast, most of the Alexa546-T4 NPs examined show quenched fluorescence (negative enhancement) compared to free Alexa 546, and the effect is only linear up to 1500 D/V (r = 0.97 for Figure 5b, red squares only). Fluorescence enhancement has been observed previously in cyanine dyes and is caused by an increase in quantum yield, which is highly dependent upon solvent polarity and viscosity.32,49,50 Upon binding to the T4, the Cy3 experiences a loss in conformational freedom, as well as a decrease in

absorbance of the viral genomic DNA inside the T4 NP and the capsid proteins, respectively, while the peaks in the visible region represent the dye absorbance. There is a small red shift (5 nm) in absorption maxima of the Cy3 upon conjugation to the T4 NP,

Figure 3. UV-vis spectra showing (a) Cy3 NHS ester (black), T4 NP after conjugation with Cy3 (red), and T4 NP (blue); (b) Alexa 546 NHS ester (black), T4 NP after conjugation with Alexa 546 (red), and T4 NP (blue). Absorbance is normalized relative to dye concentration.

Figure 4. Fluorescence emission of dye-T4 NPs. Samples were excited at 550 nm and data were normalized relative to the number of unmodified T4 NPs: (a) Cy3-T4 samples, (b) Alexa 546-samples. 599



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Figure 5. Fluorescence enhancement of dye-T4 NPs. Fluorescence enhancement of dye-T4 NPs as a function of dyes/virus calculated relative to free dye. Emissions of the Cy3-T4 and Alexa 546-T4 NPs were measured at 570 and 575 nm, respectively: (a) Cy3-T4 series; (b) Alexa 546-T4 series. Percentage of fluorescent enhancement is defined as follows: ([dye-T4 complex F.I.] - [F.I. of free dye])/[dye-T4 complex F.I.] 100.

or Alexa 546-T4 NPs (2282 D/V) at a ratio of 1 cell to 105 T4 NPs. The nuclei were visualized by DAPI staining (blue), and the dye-T4 NPs are shown as small yellow spots distributed though the cell interior. Both dye-T4 NPs, Cy3-T4 and Alexa 546-T4 NPs, were clearly visible inside cells after 4, 8, and 24 h (Figure 6a-f). Unmerged and brightfield images of Alexa-546T4 NP at 24 h incubation (Figure 6c) are shown in Figure S3 (Supporting Information). The cellular uptake was also explored by flow cytometry, which can be used to separate and quantitatively measure fluorescently labeled cell populations using different molecular probes. We measured the cellular uptake by examining the change in fluorescence between the dye-T4 treated cells and the untreated cells. Figure 7a,b shows that a clear shift is present when cells are treated with both dye-T4 NPs in comparison to untreated cells. Furthermore, the results from the flow cytometry measurements were used to quantitatively measure the uptake of the dye-T4-NPs into cells at different time points. The percentage of cells positive for dye-T4 NPs was determined by subtracting the treated histograms from the untreated. Our results indicate that the percentage of cells that are positive for dye-T4 NPs is greater than 50% at all time points, suggesting that the uptake of T4 NP is quite efficient in our treatment condition. We also used flow cytometry to determine the cell viability after cellular uptake of dye-T4 NPs by staining both treated and untreated cells with a live/dead stain. More than 92% of the cells containing the dye-T4 NPs were alive compared with >95% of untreated cells (data not shown). This suggests that dye-T4 NPs have low cytotoxicity, even after 24 h. The ability to use dye-T4 NPs for flow cytometry opens the possibility of a number of applications including cell tracking, mechanistic studies, and immunoassays. Aside from lung cancer cells, tail-less T4 NPs were also found to enter into noncancerous primary cultured cells, such as human endothelial cells, liver cells, and murine astrocytes in vitro (data not shown). While eukaryotic cells are not their natural hosts, coliphages along with their hosts are mammals’ symbiots in the intestines and some phages may pass through the membrane and circulate in the blood system. Thus, T4 may adapt to the mammalian environment and likely interacts with eukaryotic

polarity at the site of binding, both leading to an increase in quantum yield.50 Fluorescence enhancement in Alexa dyes has not been observed previously to our knowledge and may be due to energy transfer from the protein amino acids to the Alexa dye. While fluorescence enhancement is occurring in Cy3 and to a lesser extent Alexa 546 when they are bound to the T4 NP, fluorescence quenching is causing the decreases in fluorescence at higher D/V in both dyes. Quenching can arise from dyedye interactions and is often accompanied by a change in the absorption spectrum of the dye.51,52 Evidence of dye-dye interactions (such as stacking or aggregate formation) can be observed in the absorption spectra of the Cy3 labeled T4 NP in Figure 3a. The free Cy3 dye has an absorption maximum at 555 nm with a shoulder at 520 nm. When the dye is bound to the T4 NP, there is an increase in the 520 nm peak in relation to the 555 nm peak. This increase in absorbance at 520 nm or blue shift can be attributed to nonfluorescent dimer formation.50,51 A blueshifted absorption band also appears to a lesser extent in the Alexa 546-T4 absorbance spectra and has been observed in Alexa dyes previously.28,53 Dye-dye interactions are highly dependent on the location of the dyes on the virus, and while we have no information on the location of the dyes, the large surface area of the T4 NPs most likely prevents certain dye-dye quenching that would be observed with smaller VNPs. Dye-protein interactions also cause quenching by photoninduced electron transfer (PET) and have been known to play a role in the quenching of Alexa dyes.54 PET occurs when an excited dye molecule (donor) transfers its excited-state electron to another molecule (acceptor) when they are in close van der Waals contact. Recently, Chen et al. found that certain Alexa dyes were found to be quenched by interactions with tryptophan, tyrosine, methionine, and histidine residues.54 Dye-T4 NPs as Molecular Probes. The high fluorescence output and biocompatibility of dye-T4 NPs make them ideal molecular probes for cellular imaging and flow cytometry. To explore their application as molecular probes, the cellular uptake was explored using the lung cancer cell line, A549. Confocal microscopy was used to qualitatively investigate the interaction between the dye-T4 NPs and A549 cells at different incubation times (Figure 6). A549 cells were treated with Cy3-T4 (786 D/V) 600



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Figure 6. Confocal microscopy images of A549 cells after uptake of dye-T4 NPs at different time points. Uptake of Alexa 546-T4 NPs (2282 D/V): (a) 4 h after incubation, (b) 8 h, (c) 24 h. Uptake of Cy3-T4 NPs (786 D/V) after incubation at (d) 4 h, (e) 8 h, and (f) 24 h. (g) Untreated cells, negative control. Imaging was performed under 60 magnification. The scale bar is 10 μm.

Table 1. Cell Proliferation Tracking over Time timea

% positiveb

total cells

0

93

24

84.2

81 900

48

24.8

161 000

72 96

4.6 1.4

382 600 1 105 000

105 700

a Time following 6 h T4 NP treatment. b % positive for A546 T4 NP as determined by Cellometer fluorescence measurements.

Cell Tracking. The ability to fluorescently label cells and track their proliferation can be useful in a variety of in vitro and in vivo studies. To explore this application for T4 NPs, we treated A549 cells with A546-T4 (2962 D/V) for 6 h and allowed the cells to grow for up to 96 h following treatment. A cell counter equipped with a fluorescence module was used to measure the number of cells and percentage of cells positive for T4-NPs. Table 1 shows that, as the cells divided and the number of total cells increased, the percentage of cells positive for A546-T4 NP decreased (Table 1). After 96 h, the percentage of cells positive for A546T4 NPs decreased to 1.4%, a level equal to the background from untreated cells. We were also able to visualize this effect by taking images of the counted cells and observed a decrease in cellular fluorescence over time (Supporting Information Figure S4). When comparing the treated cells to untreated cells, there is no significant decrease in proliferation in dye-T4 NP treated cells (data not shown). The fluorescence of the cells was also measured by flow cytometry. Figure 8 shows that, over time, as the cells proliferate, the fluorescence of the cell population decreases and the population becomes more heterogeneous (Figure 8). This supports the hypothesis that, over time, the fluorescence signals will decrease due to the dilution of A546-T4 NPs to newly divided cells. Our results demonstrate that the internal dye-T4 can

Figure 7. Flow cytometry histograms of A549 cells treated (a) Cy3-T4 NP (786 D/V) and (b) Alexa-T4 NP (2282 D/V) at different time points after incubation, 4 h (red), 8 h (blue), and 24 h (green); untreated cells after 24 h incubation are used as negative control (black line).

cells. It is believed that the interaction with eukaryotic cells is determined by the head surface protein contents and membrane proteins. Therefore, the uptake of dye-T4 by eukaryotic cells may be specific through certain molecular interactions. Some evidence suggests that bacteriophage T4 interacts with cancer cells in vitro by binding to integrin β3 through the Lys-Gly-Asp (KGD) amino acid motif, a homologue to Arg-Gly-Asp (RGD), within the minor capsid protein, gp24.55 Likewise, our purified T4 NPs may preferentially bind and enter integrin β3þ A549 and noncancerous primary cultured cells, possibly through receptor mediated endocytosis. We are currently investigating the cellular uptake mechanisms of dye-T4 NPs. Interestingly, besides interacting with integrin β3, a nonessential head protein, Hoc, may also help to generate immune response by presenting phage T4 to macrophage and dendritic cells, through its three tandem immunoglobulin like domains.56 This suggests that macrophage and dendritic cells may also preferentially take up dye-T4 NPs. 601



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This material is available free of charge via the Internet at http:// pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]; phone: 202-404-6122; fax: 202767-9594.

Figure 8. Flow cytometric detection of cell proliferation over time following A546-T4 NP treatment.

remain inside A549 cells for at least 72 h and dye-T4 NPs can be used to quantitatively track cellular proliferation.

’ CONCLUSIONS We have successfully developed a novel viral nanoparticle, using tail-less T4 heads, as scaffolds for dye attachment, and moreover, the resulting fluorescent Cy3-T4 and Alexa 546-T4 can serve as molecular probes for cellular imaging and flow cytometry. Large surface areas and protein contents of T4 NPs give the tail-less T4 NPs more functional groups and more flexibility for dye conjugation than other viruses. The structure and size of dye-T4 NPs are close to that of unlabeled T4 NPs, and the T4 NPs are intact and stable after conjugation and purification. For the first time, we conclusively demonstrated that T4 enters eukaryotic cells using our dye-T4 NPs. Dye-T4 NPs can stay inside A549 cells at least 72 h and serve as ideal molecular probes for tracking cells, since cellular uptake of dye-T4 NPs can be clearly visualized by fluorescent microscopy, and cells positive for dye-T4 uptake can be distinguished and quantified by flow cytometry. The dye-T4 NPs can be used as a tool to understand in detail the molecular mechanisms of cell entry and the fate of the T4-NPs within cells. We are currently focusing on investigating this topic and the effect of surface modification on uptake into eukaryotic cells. Besides using dye-T4 for in vitro studies, it is also possible to use dye-T4 NPs as in vivo molecular probes, although the chemical or genetic modification strategies for more surface functionalization with target ligands and chemicals are required.6,7 The ability to conjugate the large surface area of the T4 NPs with up to 19 000 dyes/virus implies that the surface can be loaded with a variety of functionalities by using the available reactive amines. This functionalization capability will be useful in vivo when adding labels such as poly(ethylene glycol) (PEG) to prevent blood clearance, target antigens to ensure delivery to tumor tissues, and a large number of fluorescent dyes for highly fluorescent nanoprobes.

’ ACKNOWLEDGMENT We would like to thank Dr. Lindsay Black from University of Maryland for his kind gift of T4 K10. O. O. wants to thank the National Science Foundation (Grant # DMR-0648917) for financial support. We also thank Drs. James Delehanty and Jing Zhou for their review of this manuscript. This work was supported by the Office of Naval Research. The opinions and assertions contained herein are the private ones of the authors and are not to be construed as official or reflecting the views of the Department of the Navy or the military at large. ’ REFERENCES (1) Singh, P., Gonzalez, M. J., and Manchester, M. (2006) Viruses and their uses in nanotechnology. Drug Dev. Res. 67, 23–41. (2) Fischlechner, M., and Donath, E. (2007) Viruses as building blocks for materials and devices. Angew. Chem., Int. Ed. 46, 3184. (3) Mao, C., Liu, A., and Cao, B. (2009) Virus-based chemical and biological sensing. Angew. Chem., Int. Ed. 48, 6790–6810. (4) Li, K., Nguyen, H. G., Lu, X., and Wang, Q. (2010) Viruses and their potential in bioimaging and biosensing applications. Analyst 135, 21–27. (5) Lee, L. A., and Wang, Q. (2006) Adaptations of nanoscale viruses and other protein cages for medical applications. Nanomed.: Nanotechnol., Biol., Med. 2, 137–149. (6) Manchester, M., and Singh, P. (2006) Virus-based nanoparticles (VNPs): Platform technologies for diagnostic imaging. Adv. Drug Delivery Rev. 58, 1505–1522. (7) Steinmetz, N. F. (2010) Viral nanoparticles as platforms for nextgeneration therapeutics and imaging devices. Nanomed.: Nanotechnol., Biol., Med. 6, 634–641. (8) Petrenko, V. A., and Vodyanoy, V. J. (2003) Phage display for detection of biological threat agents. J. Microbiol. Methods 53, 253–262. (9) Paschke, M. (2006) Phage display systems and their application. Appl. Microbiol. Biotechnol. 70, 2–11. (10) Ackermann, H. W., DuBow, M. S., Jarvis, A. W., Jones, L. A., Krylov, V. N., Maniloff, J., Rocourt, J., Safferman, R. S., Schneider, J., Seldin, L., Sozzi, T., Stewart, P. R., Werquin, M., and Wunsche, L. (1992) The species concept and its application to tailed phages. Arch. Virol. 124, 69–82. (11) Ackermann, H. W., and Nguyen, T. M. (1983) Sewage coliphages studied by electron microscopy. Appl. Environ. Microbiol. 45, 1049–1059. (12) Ren, Z. J., Lewis, G. K., Wingfield, P. T., Locke, E. G., Steven, A. C., and Black, L. W. (1996) Phage display of intact domains at high copy number: a system based on SOC, the small outer capsid protein of bacteriophage T4. Protein Sci. 5, 1833–1843. (13) Ren, Z.-j., and Black, L. W. (1998) Phage T4 SOC and HOC display of biologically active, full length proteins on the viral capsid. Gene 215, 439–444. (14) Jiang, J., Abu-Shilbayeh, L., and Rao, V. B. (1997) Display of a PorA peptide from Neisseria meningitidis on the bacteriophage T4 capsid surface. Infect. Immunol. 65, 4770–4777. (15) Onorato, L., Stirmer, B., and Showe, M. K. (1978) Isolation and characterization of bacteriophage T4 mutant preheads. J. Virol. 27, 409–426.

’ ASSOCIATED CONTENT

bS

Supporting Information. A table containing T4 head proteins and total lysine amines; a table containing maximum fluorescence intensities for a selection of dye-T4 nanoparticles; the structure for Cy3 and Alexa 546; measurement of the number of dye/virus to the molar excess of dyes over T4 NPs; and additional cell images from confocal microscopy and Cellometer. 602



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Engineered T4 Viral Nanoparticles for Cellular Imaging and Flow

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