ABSTRACT
Enteric viruses are recognized as major etiologies of U.S. foodborne infections. These viruses are easily transmitted via food contact surfaces. Understanding virus interactions with surfaces may facilitate the development of improved means for their removal, thus reducing transmission. Using MS2 coliphage as a virus surrogate, the strength of virus adhesion to common food processing and preparation surfaces of polyvinyl chloride (PVC) and glass was assessed by atomic force microscopy (AFM) and virus recovery assays. The interaction forces of MS2 with various surfaces were measured from adhesion peaks in force-distance curves registered using a spherical bead probe preconjugated with MS2 particles. MS2 in phosphate-buffered saline (PBS) demonstrated approximately 5 times less adhesion force to glass (0.54 nN) than to PVC (2.87 nN) (P < 0.0001). This was consistent with the virus recovery data, which showed 1.4-fold fewer virus PFU recovered from PVC than from glass after identical inoculations and 24 h of cold storage. The difference in adhesion was ascribed to both intrinsic chemical characteristics and the substrate surface porosity (smooth glass versus porous PVC). Incorporating a surfactant micellar solution of sodium dodecyl sulfate (SDS) into the PBS reduced the adhesion force for PVC (∼0 nN) and consistently increased virus recovery by 19%. With direct and indirect evidence of virus adhesion, this study illustrated a two-way assessment of virus adhesion for the initial evaluation of potential means to mitigate virus adhesion to food contact surfaces.
IMPORTANCE The spread of foodborne viruses is likely associated with their adhesive nature. Virus attachment on food contact surfaces has been evaluated by quantitating virus recoveries from inoculated surfaces. This study aimed to evaluate the microenvironment in which nanometer-sized viruses interact with food contact surfaces and to compare the virus adhesion differences using AFM. The virus surrogate MS2 demonstrated less adhesion force to glass than to PVC via AFM, with the force-contributing factors including the intrinsic nature and the topography of the contact surfaces. This adhesion finding is consistent with the virus recoveries, which were determined indirectly. Greater numbers of viruses were recovered from glass than from PVC, after application at the same levels. The stronger MS2 adhesion onto PVC could be interrupted by incorporating a surfactant during the interaction between the virus and the contact surface. This study increases our understanding of the virus adhesion microenvironment and indicates ways to mitigate virus adhesion onto contact surfaces.
INTRODUCTION
Among the identifiable etiologies of U.S. foodborne diseases, a few groups of viruses are responsible for approximately one-half of the outbreak illnesses each year (1). Norovirus (NoV) is the leading causative agent for foodborne illnesses in the United States, with an estimated 5.5 million foodborne NoV illnesses each year from 2000 to 2008 (1). Critically, hepatitis A virus (HAV) is also foodborne and is known for its persistent survival in foods, e.g., produce (2), long incubation periods of up to ∼50 days after infection before symptom appearance (3), and severe disease symptoms. Via the fecal-oral route, these viruses can be transmitted through contaminated food or water or contact with contaminated surfaces. Just a few virions can cause infection in humans, who may shed virus for weeks after exposure (4).
Foodborne viruses are highly transmissible, and their spread is associated with the attaching or adhesive nature of the virions (5). While virus infection is mediated by virus adhesion via specific interactions with antigenic receptors expressed on target cells (6), pathogen adhesion on food processing surfaces may be predominantly attributed to electrostatic attraction/repulsion in combination with hydrophobic interaction (7, 8). These forces can be affected by pH, salt (ionic strength), and virus concentrations in a suspension (7). Approaches such as micromanipulation (9), atomic force microscopy (AFM) (10, 11), use of optical tweezers (12), and adsorption experiments (13) have been employed to examine bacterium and virus adhesion on various surfaces. AFM has become increasingly reliable in characterizing interaction forces in aqueous solutions with controllable environmental parameters such as the salt concentration and the presence of other compounds. A recent AFM study by Dika et al. (14) suggested that the adhesion force of virions adhering to a surface is attributable to virus and substrate interfacial charges, the ionic strength of the aqueous phase, and the hydrophobic/hydrophilic balance of both virus and substrate. The results agreed well with their virus recovery tests (14). Another AFM study of the adhesion forces of Aichi virus, MS2, and φX174 phages onto sands was conducted (11). A combined AFM and quartz crystal microbalance (QCM) approach was also used to examine the adhesion kinetics of MS2 with pristine membranes. The result revealed that the presence of a small microbial product foulant on a membrane surface inhibited virus adhesion (15). Thus, AFM may be a versatile tool to differentiate the strength of virus adhesion on various surfaces and under various environmental conditions.
The current study aimed to determine quantitatively the strength of adhesion of virions to commonly used food processing or preparation surfaces by using AFM. Similar to previous environmental studies (11, 15), MS2 coliphage was chosen as an enteric virus surrogate for this work. MS2 is similar to pathogenic enteric viruses, with a similarly sized (∼28 nm), nonenveloped, icosahedron-shaped capsid and positive-sense, single-stranded, genomic RNA (16). The disturbance of virus adhesion by a surfactant was explored for virus removal and ultimately for improvement of food safety. The surfactant sodium dodecyl sulfate (SDS) was chosen because it was proven to be effective in removing bacteria from similar substrate surfaces (17). Statistical analysis was performed to compare the adhesion forces for viruses on hydrophobic polyvinyl chloride (PVC) and hydrophilic glass. This study intended to provide direct and quantifiable values for virion adhesion using AFM, supported by indirect measurements of virus adhesion using virus recovery. Surface roughness might also play a role in virus adhesion (14). The contribution of the surface roughness of PVC to MS2 adhesion was also investigated by AFM imaging. Understanding the physicochemical forces that permit nanometer-sized viruses to attach to or to be detached from various substrates (17) and quantitatively comparing the virus recovery differences would be of benefit in developing efficient elution protocols and possible preventive controls for viral hazard entry and attachment, thereby reducing disease transmission.
RESULTS
Characterization of MS2-conjugated silicon wafer.To ensure appropriate conjugation of MS2 to the AFM tip (Fig. 1) (described in Materials and Methods), we functionalized MS2 onto a silicon wafer whose surface has chemical features similar to those of the AFM probe used. The same protocol for MS2-(3-aminopropyl)triethoxysilane (APTES) functionalization was applied. After thorough rinsing, the surface was characterized by AFM imaging. The three-dimensional (3D) image presented in Fig. 2a depicts the MS2 self-assembly on a silicon wafer via APTES cross-linking. The cross-sectional analysis illustrated in Fig. 2b showed that the maximum height of the MS2-bound wafer was within 30 nm. This is consistent with the theoretical dimensions of MS2, with a reported diameter of 24 to 26 nm, as well as the dimensions measured from our transmission electron microscopy (TEM) images of the final purified and concentrated MS2 samples used in this study (28 nm in diameter) (data not shown). It should be noted that the full width at half maximum (FWHM) of a peak of a MS2 particle measured from Fig. 2b was approximately 50 nm, which is larger than the size of a MS2 particle and might be ascribed to tip convolution (18). The result suggested that a dense layer of MS2 was conjugated successfully onto the silicon wafer via APTES cross-linking. We hypothesized that MS2 particles display similar surface features when they are functionalized on a borosilicate bead AFM probe.
Schematic presentation of MS2 conjugation onto a bare AFM tip with the APTES cross-linker.
MS2 bound to a silicon wafer surface that resembled a MS2-conjugated AFM tip surface. (a) 3D AFM image of MS2 bound to a silicon wafer surface (size of 2 by 2 by 0.058 μm3). (b) Cross section of the line scan shown in panel a. The white triangle in panel a corresponds to the dark triangle in panel b.
Measurement of adhesion forces of MS2 on PVC and glass surfaces.The adhesion forces of MS2 on glass and PVC substrate surfaces were measured using an MS2-conjugated bead probe and a bare bead probe as a control. These data are summarized in the histograms of Fig. 3a and b and were analyzed statistically. As shown in Fig. 3a, the median adhesion force of a bare tip to a glass surface was 0.19 nN, and more than 83% of the data were in the range of 0 to 0.40 nN. In contrast, the median force of an MS2-conjugated tip to a glass surface was 0.54 nN, with 80% of force readings being between 0 and 1.2 nN. Greater adhesion forces for the MS2-conjugated tip on PVC surfaces are shown in Fig. 3b, with a median force of 2.87 nN on PVC and more than 90% of the force readings being in the range of 0.6 to 8.4 nN. From the median values, the strength of MS2 adhesion to PVC was approximately 5 times higher than that to glass. Noticeably, the median force of the bare tips' adhesion to PVC was 0 to 0.4 nN, with more than 60% of the data being ∼0 nN. To understand the broad distribution of data for MS2 adhesion on PVC, we examined the surface morphology of glass and PVC surfaces in Fig. 3c and d. The z-axis represents the height of the surface feature, and the bright/dark contrast indicates various elevation regions. While the glass surface was rather smooth, the PVC surface was porous and not uniform. The smooth uniform surface of glass might be the key factor contributing to the narrow distribution of data and the small standard deviation for the forces collected. The porous surface of PVC contributes to the broad distribution of data. A large number of adhesion forces for PVC were greater than 2.87 nN, which might be ascribed to very porous regions, where stronger adhesion would be expected.
MS2 adhesion forces measured on glass and PVC surfaces. (a) Adhesion force frequency of MS2 on a glass surface, using MS2-conjugated (black bars) and bare (white bars) tips. The median force of each population is indicated by an arrow. (b) Adhesion force frequency of MS2 on a PVC surface, using MS2-conjugated (black bars) and bare (white bars) tips. The median force of each population is indicated by an arrow. (c) AFM scanning revealing a smooth glass surface, with a maximum substrate height of 1.5 nm. (d) AFM scanning revealing many pores (black dots) on a PVC surface, with a maximum substrate height of 10 nm.
Alteration of adhesion forces of MS2 on PVC with a surfactant.Understanding virus attachment and detachment was part of our purpose for this study. Once we observed the forces presented by the MS2-conjugated probe on PVC, we examined the possible force interference caused by a surfactant, SDS, during MS2 attachment to PVC. Thus, adhesion force measurements for MS2 on PVC were carried out in the presence and absence of 0.06 M SDS. As shown in the histograms in Fig. 4, SDS interfered with MS2 adhesion by reducing the median force from the original 2.87 nN without SDS to a median force of 0.021 nN with SDS in the phosphate-buffered saline (PBS), with the majority of data being distributed between 0 and 0.1 nN. Most likely, the MS2 adhesion force was disrupted and reduced by the SDS micelle layer that formed on the PVC substrate (19).
MS2 adhesion and disruptive force frequency on PVC in the presence (white bars) and absence (black bars) of 0.06 M SDS. The median force of each population is indicated by an arrow.
Recoveries from PVC and glass surfaces using a plaque assay.Because different adhesion forces for MS2 were found for the different surfaces, we determined what percentages of the virus population could be recoverable or removable from those surfaces after they were attached. With identical inoculation and elution procedures, the two substrates used in the AFM study were used for virus recovery experiments. By adjusting the inoculation volume per spot (as described in Materials and Methods), the time required for complete air drying for any inoculum on the two substrates was normalized. A large difference in drying times between the two surfaces was observed occasionally, but the difference was still less than 5 min across all five trials, with a drying time of approximately 30 min for each sample on either surface per trial.
Our early data indicated that insignificantly different quantities of viruses were recovered from the two surfaces 1 min after inoculation (data not shown). Other types of surfaces were tested similarly and showed similar recoveries, compared with the glass and PVC surfaces. This may be due to the lack of firm attachment after only 1 min of contact time. Thus, a period of 24 h of cold storage after drying was used to allow stronger virus attachment to the substrate surfaces. After complete air drying and 24 h of cold storage, the amount of recoverable MS2 from glass was always greater than that from PVC, with recovery ratios of 1.32 to 1.50 (Table 1). Additional analysis of variance (ANOVA) of all virus recoveries in the five trials indicated that recoveries from the glass substrates were significantly (P = 0.002) greater than those from the PVC substrates, with 2 to 4 samples of each substrate type in each trial. These recoveries indicated that the population of unrecoverable (or firmly attached) MS2 on PVC remained greater than that on glass after 30 min of air drying and 24 h of cold storage. The lower MS2 recoveries from PVC, according to infectivity, seemed to be in agreement with stronger adhesion forces for MS2 on PVC than on glass substrates, as determined by AFM.
Virus recoveries from inoculated glass and PVC surfaces after 24 h of cold incubation, without SDS treatment
Since the MS2 adhesion force was disturbed by the presence of SDS during the AFM studies (Fig. 4), virus recovery from SDS-treated PVC was also evaluated. Our first observation was that the MS2 inoculum droplet containing SDS spread out as if on a glass surface, indicating that the interaction of MS2 with the hydrophobic PVC surface had been altered. It was also found that the viability of the MS2 suspension incubated with SDS did not change after storage at 5°C for 4 h. In order to compare the recoveries between the surfaces with and without SDS and to maintain similar drying times for the two surfaces in trial 601-2, the PVC surfaces with SDS and without SDS were spotted with 1 drop of 10 μl of inoculum and 2 drops of 5 μl of inoculum, respectively. The results showed an increase in recovery from the SDS-treated PVC surface of 19%, compared to the nontreated PVC surface (Table 2). The increased recovery was in agreement with the disrupted/weakened adhesion force for MS2 on PVC with SDS that was seen using AFM.
Virus recoveries from inoculated PVC surfaces after 4 h of cold incubation, with or without SDS treatment
DISCUSSION
The Centers for Disease Control and Prevention estimate that, each year, one in six Americans acquire a foodborne illness, with viral infection being the most likely cause (1, 20). To reduce or to prevent foodborne viral infections via spread to and from food contact surfaces, an understanding of the factors affecting virus attachment to surfaces is crucial. This understanding could suggest means to interrupt transmission of viruses and aid removal of viruses from these surfaces. In previous studies on virus attachment, adhesion, or transmission on specific surfaces or food, scientists frequently evaluated virus recoveries from inoculated surfaces through elution and quantitation using plaque assays (5) or PCR methods (21). These bulk recovery methods do not allow an understanding of the force interactions between the virus and the surface. The current study aimed to evaluate the surface microenvironment and the interactions of the virus with the substrates by quantitatively assessing the adhesion strength of virions with food contact surfaces using AFM and comparing the results to those obtained using a standard elution and virus quantitation assay. Therefore, the strength of virus adhesion was measured directly by AFM and indirectly by virus recovery.
Initial experiments indicated the presence of surface-active materials in the virus stock, which were attributed to the host cell growth medium and host cell fragments. These compounds, which potentially interfered with AFM readings, were removed through inclusion of a foam fractionation technique; this resulted in purified virus stock solutions, as confirmed by dynamic light scattering (DLS), TEM, and surface tension measurements. To prove that this cleaning protocol was appropriate for the AFM requirements, the host bacterial cell solution (without viruses) was also subjected to foam fractionation and evaluation and was found to demonstrate negligible adhesion forces (data not shown). In contrast to the MS2-conjugated AFM tip, nonconjugated bare AFM tips in the study showed forces of close to 0 nN for both PVC and glass surfaces (Fig. 3a and b). With negligible signals imparted by the multiple controls, all AFM force readings should be attributed to virus-surface interactions.
MS2 monolayer conjugation to an AFM tip was evaluated using an APTES-functionalized silicon wafer as a model system (see Results). Via silane coupling (22), MS2 virions were conjugated by the interaction of the amine at the end of APTES and the carboxylic acid of the MS2 capsid proteins. The APTES concentration was optimized at 4% after comparing three-phase contact angles of water droplets on the MS2-conjugated wafers (23) at various APTES concentrations (1% to 8%) (data not shown). Monolayer coverage was confirmed by the uniform dense distribution of MS2 across the wafer surface (Fig. 2b). Our preliminary work using sharp AFM tips (∼30 nm at the tip apex) generated lower adhesion forces due to the smaller contact area. This led to lower signal strength than that for the background. In this work, 5-μm spherical AFM probes were used. Spherical tips provide a larger surface to conjugate more virions, for more reliable measurements (15).
Previously, Attinti et al. (11) reported a MS2 adhesion force of 0.49 nN with oxide-removed sands (hydrophilic, with a surface charge of −39.5 mV), compared to a force of 19 nN with aluminum oxide-coated sands (hydrophobic, with a surface charge of +4.1 mV). This force difference was possibly due to more electrostatic attraction and fewer hydrophobic interactions (11). In the current study, an adhesion force of 0.54 nN was found with glass, which is relatively hydrophilic and negatively charged (24). These two characteristics of glass are similar to those found for the oxide-removed sands (11), which may be a reason why MS2 demonstrated similar adhesion forces with our glass surface and those authors' oxide-removed sands. The adhesion forces of MS2 with PVC were greater (2.87 nN), although not as great as those with oxide-coated sands (11). PVC is hydrophobic and negatively charged (24). Although the lesser forces on PVC, compared with aluminum oxide-coated sands, may be due to electrostatic interactions, other factors may play a role. The factors that could logically affect the adhesion force readings include the virion contact area, the AFM probe shape and size, and substrate surface asperities (14, 22, 25). Therefore, the AFM force readings for different substrates should not be considered absolute values. For instance, data fluctuations were observed when we measured MS2 adhesion to the same substrate with different probes (data not shown). This finding was mainly due to different probes having different contact surfaces that permitted different amounts of MS2 to be conjugated at the contact regions. However, the relative force differences (for example, the median glass/PVC force ratio) should remain approximately the same. Using the same MS2-conjugated probe to measure adhesion to glass and to PVC, as we did in the current study, validates direct comparisons of the force values. In addition, the broader distribution of the force data for MS2 with the PVC substrate (Fig. 3b) in the current study was possibly due to the porous nature of the surface. Dika et al. (14) indicated that increased roughness of a surface increases the overall surface area available for virus adhesion or attachment. According to Katainen et al. (26), for interactions between a particle and a single substrate asperity, the strength of the adhesion is largely determined by the contact area between the particle and the asperity on the substrate. In the current study, the contact area between the MS2-conjugated AFM tip and the PVC surface would be more variable and inconsistent than that for the glass substrate, possibly due to larger numbers and increased sizes of asperities on PVC, as illustrated by the surface images in Fig. 3c and d. Therefore, the median values of the measured adhesion forces were used to represent the strength of MS2 adhesion to each substrate. In fact, the roughest areas of PVC resulted in MS2 adhesion forces more than 5 times greater than those for glass, as the average reported here (ranging from 7 times to 3 times greater in different trials). The AFM adhesion force results for MS2 with the two substrates were supported by the MS2 virus recoveries (Table 1). Stronger adhesion or interaction of the PVC substrate with MS2 resulted in lower recoveries, compared to weaker adhesion with the glass substrate, with an average PVC/glass virus recovery ratio of 1:1.4 after MS2 incubation on the surfaces for 1 day at 5°C.
Understanding the physicochemical forces that allow viruses to attach to or be disrupted from attaching to various substrates and quantitatively comparing their affinities for attachment to different food contact surfaces would be of benefit in developing preventive controls for surfaces, thereby reducing disease transmission. In this study, MS2 adhesion forces for PVC were disrupted by the inclusion of SDS, presumably due to the micellar layer's repulsive force from the micellar dispersion (19, 27). SDS was found to have no significant lytic activity for some Gram-negative organisms (28), and many bacteria can even grow in the presence of 5% SDS (29) at neutral pH. A previous study showed that treatment of murine norovirus (MNV) with SDS at 10,000 ppm at 37°C caused a reduction of several log units after 72 h of incubation and a reduction of <1 log unit after 4 h of incubation (30). Our pretests showed negligible MS2 inactivation using 0.06 M SDS (equivalent to 17,000 ppm SDS) for up to 4 h at room temperature. Therefore, we think that the MS2 viruses mixed with 0.06 M SDS were still alive during the 4-h examination by AFM at room temperature during each trial of the current study. Recently, a study showed that 0.5% SDS combined with 0.5% levulinic acid solution improved virus removal from strawberries (31), with 2.4 and 2.7 log units of MS2 and HAV, respectively, being removed, compared to an average of only 1.4 log units being removed by water washing alone. In the current recovery study, SDS solution was applied first, before inoculation, to prevent or to interfere with virus attachment to abiotic surfaces. When a solid surface is first exposed to the SDS solution (0.06 M, i.e., 1.7% or 17,000 ppm), it is likely that a simple adsorbed surfactant monolayer or an SDS micellar layer is formed on the surface (19), which prevents any subsequently introduced virions from directly contacting the surface. Therefore, the use of an aqueous solution containing SDS to coat food contact surfaces may provide a means of preventing cross-contamination by disrupting the adhesion forces between the virions and the contact surface, as shown in Table 2. The differences in virus adhesion forces between glass and PVC were confirmed by indirectly measuring virus recoveries from the two surfaces. The lower MS2 recoveries from PVC, according to infectivity, seemed to be in agreement with the stronger adhesion force found for MS2 with PVC, compared with that for glass, as determined by AFM.
In summary, in past studies looking at specific surfaces, virus attachment or adhesion was evaluated by quantitating recoveries from inoculated surfaces via infectivity assays or molecular assays (e.g., reverse transcription [RT]-PCR). In addition to phenotypic virus recovery determination, this study aimed to evaluate the microenvironment in which nanoparticles (virions) interacted with select food contact surfaces, by quantitatively comparing the adhesion strength of virions using AFM. Our data indicated that the median MS2 adhesion force with PVC was approximately 5 times greater than that with glass. Our virus recovery studies indicated that PVC surfaces allowed less bulk recovery, with a PVC/glass recovery ratio of 1:1.4, indicating that the MS2 was more strongly attached to PVC than to glass. The difference in MS2 adhesion to these two substrates was ascribed not only to the intrinsic nature of the substrate (glass or PVC) but also to the substrate topography (smooth or rough). Additionally, we proved that the stronger MS2 adhesion forces for PVC could be weakened by incorporating a micellar solution of a surfactant such as SDS. This study increases our understanding of the microenvironmental determinants of virus attachment, perhaps facilitating rapid screening of potential means to mitigate virus adhesion to food contact surfaces.
MATERIALS AND METHODS
MS2 stock preparation and purification.MS2 coliphage stock was propagated and enumerated using the double agar layer (DAL) plaque assay (32), with Escherichia coli (ATCC 700891) as the host. After propagation, E. coli host cell debris was removed by centrifugation at 11,000 × g for 20 min. For further purification, foam fractionation was completed using the Bartsch method (33). Briefly, the MS2 in spent growth medium was shaken briskly in a Pyrex bottle to produce a foam layer, which captured impurities and E. coli host cell debris. The foam was removed by vacuum via a Pasteur pipette, without touching the liquid layer. This method was repeated several times until foam production was significantly reduced. The remaining suspension was filtered through Whatman 0.1-μm-pore-size track-etched polycarbonate membranes (GE Healthcare, Pittsburgh, PA). Additional purification to remove small particulates (<100-kDa) from the virus suspension was carried out by ultrafiltration using a 100-kDa Amicon unit (EMD Millipore, Billerica, MA), with a final rinse with PBS (tissue culture grade). The purified and concentrated MS2 stock was sterile-filtered again using a 0.22-μm-pore-size filter and then stored at 5°C. This final virus sample solution was approximately 108 to 109 PFU/ml, as determined with the DAL plaque assay.
To ensure the virus sample quality, we carried out equilibrium surface tension measurements using a K8 Wilhelmy plate tensiometer (Krüss GmbH, Hamburg, Germany). We repeated the following process for measuring the surface tension of virus (MS2) stock solutions. Twenty milliliters of deionized (DI) water was poured into a petri dish, the end of the platinum plate of the equipment was vertically touched to the top surface of the water, and then the plate was lifted by applying a small amount of force slowly, indicating the surface tension (34). At 25 ± 2°C, the final surface tension increased to 60 mN/m after the foam fractionation was applied to the sample, from the original reading of ∼45 mN/m without foam fractionation. This indicated that, via the foam fractionation, there was removal of surface-active impurities such as phospholipids and debris from the E. coli host cells. Furthermore, the DLS technique (also called photon correlation spectroscopy [35]), which is a technique to monitor the distribution of nanometer-to-micron-sized particles in bulk solution, was used to determine the particle size distribution in solutions, using a Zetasizer 3000 HSA sizer (Malvern Instruments, Malvern, UK). The average hydrodynamic diameter of the MS2 virions (i.e., 28 to 34 nm [15]) was 32 ± 3 nm, when measured in PBS (pH 7.4.) at 25 ± 2°C.
Preparation of MS2-functionalized AFM probes and adhesion force measurements.Silicon nitride AFM cantilevers with a 5-μm-diameter borosilicate glass bead probe (Novascan, Ames, IA) were used in the study. Prior to AFM measurements, the probe was cleaned by soaking the cantilever in H2SO4/H2O2 (7:3 [vol/vol]; Sigma, St. Louis, MO) for 5 min at room temperature. After rinsing with DI water, the cantilever was boiled in DI water for 10 min. Similar to published protocols (36, 37), MS2 was functionalized on AFM probes in this study. As shown in Fig. 1, the cantilever was incubated in 4% (vol/vol) APTES (Sigma-Aldrich) in toluene (Sigma-Aldrich) for 3 h at 60°C, followed by washing twice with toluene and once with ethanol (200 proof; Sigma) to remove residual APTES. The APTES-treated cantilevers were dried at 60°C for 5 min before soaking overnight at 5°C in a suspension of ∼108 PFU/ml MS2 in PBS, prepared as described above. The amino groups within APTES permitted conjugation between virions and the AFM probe. A control (bare) probe was also prepared with no APTES treatment; thus, it is unlikely that MS2 was conjugated on the control tip.
The adhesion force measurements were carried out using an Agilent 5500 atomic force microscope (Agilent Technologies., Santa Clara, CA). The force-distance curves were recorded in fluid contact mode with an MS2-conjugated probe, in PBS at pH 7.4. The z-scan rate was 0.5 μm/s. The spring constant was calibrated by using reference cantilevers with known spring constants. The spring constants were 0.0103 ± 0.006 and 0.0173 ± 0.009 N/m for an APTES-untreated (bare) probe and an MS2 conjugated probe, respectively. Typically, 16 force-distance curves were collected in areas of 1 μm by 1 μm or 64 curves were collected in areas of 15 μm by 15 μm at each location; nine locations were chosen in each trial. Three independent trials were conducted. More than 100 data points were collected and analyzed statistically via ANOVA for PVC surfaces (catalog no. s17525B; Fisher Scientific, Pittsburgh, PA) and glass surfaces (catalog no. 12-550-A3; Fisher Scientific). The maximum downward force exerted on the cantilever in retraction curves is referred to as the adhesion force (measured relative to the baseline), and values were used to build the adhesion force histograms (10). To evaluate whether MS2 virion attachment could be disrupted, the adhesion force measurements were also conducted in the presence of 0.06 M SDS (Sigma-Aldrich) micellar solution. SDS was chosen because it was proven to be effective in removing bacteria from the same substrate (17). The concentration of 0.06 M SDS was used because it is several times higher than the critical micelle concentration (CMC). At this concentration higher than the CMC, the SDS micellar layer preformed on PVC surfaces exhibits a repulsive force (19, 27) to MS2 attachment on the PVC.
In order to ensure that MS2 virions were coated properly on the AFM probe, silicon wafers (Cabot Microelectronics, Aurora, IL) were functionalized with MS2 by following the same protocol as described above. The MS2-silica surface was examined using an Asylum MFP-3D atomic force microscope in fluid tapping mode, to confirm the MS2 coliphage conjugation on the APTES-treated silicon wafer surface. A sharp silicon nitride AFM tip (BudgetSensors, Sofia, Bulgaria) was used to scan the surface in PBS at pH 7.4. The tip radius was approximately 15 nm, the cantilever length was 200 μm, and the spring constant was 0.04 ± 0.01 N/m. The scanning area was 2 μm by 2 μm, the scanning rate was 0.5 Hz, and the scanning speed was 2.5 μm/s, with a driving frequency of 8.5 kHz (38). Gwyddion software (Brno, Czech Republic) was used to analyze the scanned images in two and three dimensions.
MS2 recoveries from glass and PVC surfaces.PVC and glass substrate surfaces were extensively cleaned with a laboratory detergent (Sparkleen 1; Fisher), using gloved-hand massage and a soft brush, followed by two DI water rinses by shaking at 100 rpm for 30 min each at room temperature. Finally, the substrates were rinsed with 70% ethanol using the same rinse protocol of 100 rpm for 30 min. The substrates were transferred aseptically to a biosafety cabinet, using sterile tweezers, for air drying prior to virus inoculation.
Each substrate was inoculated with the same concentration of virus; however, to adjust for the different drying times caused by the hydrophilic and hydrophobic surfaces, different volumes (or droplet numbers) for the same total inocula were inoculated onto the two surfaces. This allowed the inocula to be completely dry within the same time frame, which minimized the variability that could be contributed by different drying times. Glass slides were inoculated with one spot of 10 μl and PVC surfaces were inoculated with two spots of 5 μl of identical MS2 solutions and concentrations. Slides were dried for approximately 30 min and stored at 5°C for 24 h prior to virus recovery, to allow sufficient time for virus attachment. For virus recovery, MS2 was eluted from the slides using 5 ml of eluent containing 1% beef extract with 50 mM glycine and 100 mM Tris (pH 8), in a 50-ml conical tube at a 45° angle, with shaking at 100 rpm for 10 min. Positive controls were prepared by direct addition of the inoculum to the elution tubes. The MS2 DAL plaque assay (17) was used to enumerate the recovered phage, and recoveries were calculated based on the positive-control tubes.
To determine any changes in virus recovery caused by disrupted/weakened virus adhesion, PVC surfaces were treated with a surfactant, SDS, before virus inoculation. Specifically, 10 μl of 0.06 M SDS in PBS was inoculated onto each PVC surface and partially dried (until a ring stain formed, approximately 15 min) before the surface was inoculated with 10 μl of MS2 on the same spot. Control samples without SDS were also prepared by inoculating 1 spot of 10 μl per surface or 2 spots of 5 μl per surface. After complete air drying, inoculated PVC surfaces (with or without SDS treatment) were stored at 5°C for 4 h, and then viruses were eluted for the recovery comparison. Earlier experiments showed that 4 h of incubation with SDS did not affect MS2 survival.
ACKNOWLEDGMENTS
We thank FDA staff members A. Shazer and K. Reineke for their assistance. We appreciate M. L. Tortorello and R. E. McDonald for their comments on the manuscript.
This research would not have been possible without funding from U.S. Food and Drug Administration.
FOOTNOTES
- Received 31 August 2017.
- Accepted 2 October 2017.
- Accepted manuscript posted online 6 October 2017.
- This is a work of the U.S. Government and is not subject to copyright protection in the United States. Foreign copyrights may apply.
