Surface-Templated Nanobubbles Protect Proteins from Surface

Letter Cite This: J. Phys. Chem. Lett. 2019, 10, 2641−2647

pubs.acs.org/JPCL

Surface-Templated Nanobubbles Protect Proteins from SurfaceMediated Denaturation David S. Bull, Daniel F. Kienle, Andres F. Chaparro Sosa, Nathaniel Nelson, Shambojit Roy, Jennifer N. Cha, Daniel K. Schwartz, Joel L. Kaar, and Andrew P. Goodwin* Department of Chemical and Biological Engineering, University of Colorado Boulder, Boulder, Colorado 80303, United States

J. Phys. Chem. Lett. Downloaded from pubs.acs.org by WESTERN UNIV on 05/09/19. For personal use only.

S Supporting Information *

ABSTRACT: In this Letter, we report that surface-bound nanobubbles reduce protein denaturation on methylated glass by irreversible protein shell formation. Single-molecule total internal reflection fluorescence (SM-TIRF) microscopy was combined with intramolecular Förster resonance energy transfer (FRET) to study the conformational dynamics of nitroreductase (NfsB) on nanobubble-laden methylated glass surfaces, using reflection brightfield microscopy to register nanobubble locations with NfsB adsorption. First, NfsB adsorbed irreversibly to nanobubbles with no apparent desorption after 5 h. Moreover, virtually all (96%) of the NfsB molecules that interacted with nanobubbles remained folded, whereas less than 50% of NfsB molecules remained folded in the absence of nanobubbles on unmodified silica or methylated glass surfaces. This trend was confirmed by ensemble-average fluorometer TIRF experiments. We hypothesize that nanobubbles reduce protein damage by passivating strongly denaturing topographical surface defects. Thus, nanobubble stabilization on surfaces may have important implications for antifouling surfaces and improving therapeutic protein storage.

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Here, we show that surface-bound nanobubbles promote protein stability on hydrophobically modified glass surfaces because of their specific deposition at topographical or chemical surface defect sites. Nanobubbles are small, stable gas pockets that may be present at a solid−liquid interface in air-saturated solutions, with dimensions on the order of tens of nanometers in height and hundreds or thousands of nanometers in diameter. 16−19 These nanobubbles have been experimentally confirmed through multiple characterization methods, including atomic force microscopy,16,20 attenuated total reflection infrared spectroscopy,21 total internal reflection fluorescence microscopy (TIRF),22 and others.23−28 Surface nanobubbles are nucleated from a supersaturated gas solution at a liquid−solid interface either through direct immersion of a surface in water20 or solvent exchange from a solvent with higher gas solubility.29 Nanobubble nucleation is followed by growth from dissolved gas controlled by either gas saturation or temperature.30−32 One prominent theory is that nanobubbles obtain their exceptional stability from pinning of the nanobubble contact line at both structural and chemical heterogeneous surface sites, the same sites theorized to promote protein denaturation.18,33−36 As such, most nanobubble studies have been reported on hydrophobic surfaces that contain inherent defects such as step edges, which are the same anomalous sites that contribute to surface-mediated protein unfolding.35−39 Furthermore, studies have shown that

roteins are ubiquitous in biomedical and biotechnological applications, including protein therapeutics, biosensing, biocatalysis, food processing, and detergents.1−3 Unfortunately, proteins are notoriously unstable over long-term storage as a result of air oxidation, thermal unfolding, mechanical shearing by ice crystals, and many other potential processes.4 Beyond simple loss of activity, in some cases therapeutic proteins form large aggregates that can provoke an immune response, diminishing long-term efficacy and potentially even endangering the patient’s life.5−9 Thus, a considerable amount of research and effort has been devoted to maintaining protein stability during storage.6,10,11 Even under optimal conditions, proteins still adsorb and desorb from surfaces that contact liquid formulations, in some cases desorbing from the surface in an unfolded state. Recent advances using single-molecule (SM) fluorescence microscopy methods have led to the identification of anomalous surface sites, or “hot spots,” that are responsible for the majority of unfolding events.12 These anomalous denaturing sites are thought to be caused by random heterogeneities in the surface (e.g., glass) structure such as step edges or grain boundaries. Even nominally smooth glass exhibits topographical nonuniformities with protruding or depressed surface features, which can, in turn, alter the interaction of the protein with the surface relative to homogeneous surfaces.12−14 Because these features may arise from many sources, such as demixing, crystallization, contamination, and/or the processing history of the glass,15 and are thus generally unavoidable, an understanding of these sites and strategies to mediate their impact on protein stability are critical. © XXXX American Chemical Society

Received: March 21, 2019 Accepted: May 6, 2019

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DOI: 10.1021/acs.jpclett.9b00806 J. Phys. Chem. Lett. 2019, 10, 2641−2647

Letter

The Journal of Physical Chemistry Letters nanobubbles interact with protein, especially at the three-phase line, and thus a study on the effect of nanobubble coated surfaces on protein confirmation is relevant.40,41 By comparing the conformation of proteins resulting from interactions with nanobubbles, methylated glass, and unmodified silica surfaces, it was found that the presence of nanobubbles imparted far greater conformational stability to proteins than a surface in the absence of nanobubbles. We show that such nanobubbles mitigated the denaturing effects of these defect sites through formation of a sacrificial, protective protein adlayer on the nanobubble surface. These observations are likely to provide useful information not only for stabilizing proteins in storage but also in biosensing applications and in the immobilization of enzymes for biocatalysis. In order to observe specific protein interactions with surface nanobubbles, SM trajectories of individual dye-labeled protein molecules were tracked by TIRF microscopy. The use of TIRF limits fluorescence excitation to a region extending only ∼100 nm from the surface, thereby allowing selective tracking of proteins during their interaction with the surface in an otherwise high fluorescence background. As compared to more conventional, ensemble protein characterization methods, SM experiments are able to distinguish distinct dynamic behaviors of individual proteins, which in turn may be analyzed statistically to predict bulk behavior. This technique has been utilized to study protein adhesion, diffusion modes, and conformational changes as a result of protein−surface interactions.12,42−44 Here, the enzyme nitroreductase (NfsB) was used as a model protein for tracking protein interactions with surface nanobubbles. We have previously labeled NfsB with dyes capable of Förster resonance energy transfer (FRET) and showed that the labeled construct could be used to distinguish folded from unfolded states in SM studies.42 In addition, NfsB has been extensively studied given its utility to neutralize explosives among other applications, which potentially require the enzyme to be stable during storage over long times.42,45−47 Two NfsB constructs were used in these studies: the first construct was singly labeled with AlexaFluor 488 (AF488-NfsB) to track protein exchange on surfaces, and the second construct was dually labeled with AlexaFluor 555 and CF633 (AF555-CF633-NfsB) to enable FRET.42 First, the general behavior of NfsB on nanobubbles was determined. To form nanobubbles, #0 glass slides were methylated by chemical vapor deposition of hexamethyldisilazane. The macroscopic uniformity of the trimethylsilyl (TMS) coating was confirmed by the observation of an 88 ± 3° contact angle in three separate locations as measured by sessile drop goniometry. Advancing and receding contact angles were taken yielding 100.4 ± 1° and 85.6 ± 1°, respectively, over five different TMS-coated silica surfaces. Nanobubbles were deposited onto these surfaces using an ethanol−water solvent exchange method described previously (Figure S1).48 After nanobubble deposition on methylated glass by solvent exchange, AF488-NfsB molecules were found to stick onto nanobubble sites as shown by TIRF microscopy (Figure 1A). The high intensity of the TIRF excitation laser caused rapid photobleaching of the nanobubble-adsorbed AF488-NfsB (Figure 1B). In order to observe if the NfsB desorbed from the nanobubble surface, fluorescence recovery after photobleaching (FRAP) was used. The AF488-NfsB bound to the nanobubbles was photobleached, and the rate of exchange with protein in the bulk solution was followed by periodically capturing time-lapse images (Figure S2). However, no

Figure 1. (A,B) Fluorescence images of AF488-NfsB adsorbed to nanobubbles at (A) 0 s and (B) 60 s of continuous irradiation, showing photobleaching of AF488-NfsB. (C) Normalized emission intensity of AF488-NfsB fluorescence monitored for 5 h showing the absence of recovery associate with solution-phase exchange, with photobleaching finishing at t = 0. To avoid bleaching during recovery phase (t > 0), images to construct graph (C) were acquired every 5 min. The shaded region shows intensity from images acquired prior to photobleaching.

fluorescence recovery was measured even after 5 h (Figure 1C), suggesting that virtually no exchange occurred between adsorbed and bulk protein molecules. Thus, proteins appeared to form a stable adlayer on the nanobubble surface that remained for a long period of time. This finding is consistent with the results of other studies that show proteins adhere strongly to the air−water interface.49 Following the bulk protein adsorption studies described above, the interaction of individual AF555-CF633-NfsB molecules were observed with nanobubbles using SM-TIRF. In order to determine how proteins interacted with nanobubbles, it was necessary to develop an imaging method to define the spatial location of each nanobubble site. Because of the large difference in refractive index between air and water, we hypothesized that reflection bright field microscopy could resolve nanobubbles without using a fluorescence channel, potentially making this approach orthogonal to SM-TIRF. As shown in Figure 2A, nanobubbles appeared as dark spots on a gray background, even without a fluorescent marker to highlight the bubbles. From these images, the nanobubbles were identified by edge detection (MATLAB, Mathworks Inc.), shown as circles in Figure 2A. Thus, for each molecule of AF555-CF633-NfsB that was observed in SM-TIRF images, the molecule was assigned to a specific nanobubble site or the surrounding TMS-glass surface (Figure 2B). The donor and acceptor emission intensities of each AF555CF633-NfsB molecule were measured in each protein trajectory that interacted with the nanobubble-adlayer surface (Figure S3). To characterize transient protein interactions at 2642


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trajectories at the same spot, tens of thousands of trajectories (see Table S1) for each surface were accumulated. Examples of colocalization between the NfsB trajectories (SM-FRET) and nanobubbles (reflection bright field) are shown in Figure 3A. The folded and unfolded conformational states could be clearly distinguished by comparing the ratio of donor (FD) and acceptor (FA) emission intensities (Figure 3B). To quantitatively distinguish conformational states, a threshold line was assigned at the saddle point between the two peaks in the heat map, which minimized the integrated values of the heat map along the line.42 Each time step of a trajectory was categorized as folded or unfolded based on its position relative to the threshold line. In some cases, changes in folding state were observed within a given trajectory, which was indicated by the change in position of the location of the donor and acceptor intensities relative to the threshold line in the heat map (Figure 3). To determine the effect of nanobubbles on NfsB conformation, methylated surfaces were prepared by exchanging ethanol with air-saturated water (with nanobubbles) or argon-degassed water (without nanobubbles). Reflection bright field imaging was used to confirm the presence or absence of nanobubbles. When a 10−12 M solution of AF555CF633-NfsB was exposed to a nanobubble-laden surface, 74 ± 3% of the proteins were folded upon leaving the surface (Table S1). For the control surface without nanobubbles, only 42 ± 4% of the protein remained folded upon desorbing from the methylated surface, and only 54 ± 1% of the protein remained folded after interaction with unmodified silica. To study the effect of protein concentration, the 10−12 M labeled protein was added to either 10−10, 10−8, or 10−4 M unlabeled NfsB to allow counting of SM trajectories. Increasing the solution protein concentration also increased the fraction of events in which the protein desorbed in a folded state (Figure 4): 10−10 M solution concentration yielded an 87 ± 1% folded fraction, 10−8 M yielded an 86 ± 2% folded fraction, and 10−4 M NfsB yielded a 96 ± 1% folded fraction. Remarkably, the opposite

Figure 2. (A) Reflection bright field imaging of nanobubbles, with nanobubble regions selected. (B) Representative image of laser illuminated AF555-CF633-NfsB with overlaid nanobubble regions.

nanobubble sites, AF555-CF633-NfsB was introduced to the surface and illuminated by alternating-laser TIRF excitation. Upon illumination, the donor and acceptor fluorescence intensity emitted from AF555-CF633-NfsB was acquired in spectrally distinct optical channels that were spatially aligned, using alternating laser excitation to only collect data with both donor and acceptor emission. This additional data filtering step prevented potential artifacts due to mislabeling and photobleaching. Using a low concentration of fluorescently labeled NfsB (10−12 M) to prevent simultaneous detection of multiple SM

Figure 3. (A) SM trajectories (colored lines) of AF555-CF633-NfsB colocalized on reflection bright field microscopy images of nanobubbles. Each superimposed line represents a different trajectory with colors added to help differentiate trajectories. (B,C) Example trajectories of a single NfsB molecule’s interaction with either (B) a glass surface or (C) a nanobubble containing surface, demonstrating the dynamic nature of protein interaction with the surface in changes of FD and FA over time. Red shading indicates low FRET (unfolded protein) and blue shading indicates high FRET (folded protein). (D,E) Corresponding FRET trajectories are overlaid onto a heat map of all trajectories interacted on this surface, where B and D display a trajectory from silica with 10−4 M protein concentration while C and E display a trajectory from 10−12 M protein concentration on a nanobubble surface. For clarity, only initial, final, and folding/unfolding transitions have been mapped. 2643


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nanobubbles and the NfsB adlayer) or if the interaction actively caused refolding. Because of the limited temporal resolution of these experiments (70 ms acquisition time), it was difficult for the SM TIRF microscopy studies alone to differentiate between these states if the refolding occurred quickly. In order to distinguish between these possibilities, NfsB was thermally denatured prior to exposure to the surfaces and the extent of refolding of NfsB was quantified. Enzymatic activity assays (Figure S4) showed that heating the NfsB to 55−60 °C resulted in the loss of activity, which was irreversible upon cooling to room temperature. Irreversible denaturing was also confirmed by circular dichroism upon heating the protein from 20 to 80 °C; again, the protein did not return to its initial state upon cooling to room temperature (Figure S4). Based on these results, AF555-CF633-NfsB was denatured prior to SM experiments by heating the protein to 65 °C for 15 min. SM TIRFM-FRET experiments were performed using 10−12 M denatured AF555-CF633-NfsB and 10−4 M denatured NfsB in deionized water. As evident by a heat map of accumulated raw trajectories, the initially denatured protein was observed to exhibit a broad range of folding states, neither predominately folded or unfolded (3001 trajectories), upon desorption from nanobubble-laden surfaces. Initially denatured NfsB desorbing from a bare methylated surface without nanobubbles showed a very similar diffuse heat map (1748 trajectories) (Figure 5). Since circular dichroism studies showed that the protein remained unfolded in solution once heated (Figure S4), these diffuse heat maps (depicting extremely broad distributions of FRET signals) indicated that neither surface actively promoted protein refolding. In particular, refolding of the protein would

Figure 4. (A) Heat maps showing donor fluorescence and acceptor fluorescence for NfsB on various surfaces (top to bottom): Silica, unfunctionalized glass; −NB, methylated without deposited nanobubbles; +NB, methylated without deposited nanobubbles. Each dashed line represents the division between unfolded (left of line) and folded (right of line). Each diamond represents the peak position of the unfolded protein population, and each star represents the peak position of the respective folded protein population. (B) Fraction of folded NfsB on surfaces vs bulk NfsB concentration for each data set in A. Blue circles, TMS-coated surfaces with deposited nanobubbles; red squares, TMS-coated surfaces without deposited nanobubbles; black diamonds, unmodified clean silica surfaces without deposited nanobubbles. Error bars are one standard error as estimated by jackknife resampling; in some cases, the error bar is smaller than the data point.

trend was found for surfaces without nanobubbles. In the case of the methylated glass without nanobubbles, only 29 ± 7% of the protein molecules left the surface while folded, while only 22 ± 2% of the protein remained folded on unmodified glass. Thus, the presence of nanobubbles increased the fraction of desorption events in which the proteins remained folded, when compared to both methylated glass and unmodified silica surfaces; at higher concentrations, unfolding was eliminated almost entirely. Next, we sought to determine if the high folded fraction of NfsB was due to the inhibition of NfsB unfolding (mediated by

Figure 5. (A) Heat maps of donor fluorescence and acceptor fluorescence. (B) Normalized fluorescence emission spectra obtained in flow cells for intact or denatured AF488-CF633-NfsB (AFCNN) on TMS-coated surfaces with or without nanobubbles. Blue: intact, +NBs; red: intact, −NBs; green: denatured, +NBs; purple: denatured, −NBs. Inset: acceptor emission region. 2644



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have resulted in a peak of high intensity in the area associated with the folded state in Figure 3. Overall, these results implied that the mechanism of stabilization of AF555-CF633-NfsB by the nanobubbles was due to the protective effect of the nanobubbles, rather than refolding of AF555-CF633-NfsB by the nanobubbles. Finally, the SM observations were connected with bulk macroscopic measurements of protein stability on the surfaces of interest. This was addressed by measuring the ensemble conformation of AF555-CF633-NfsB in contact with nanobubble-laden and nanobubble-free surfaces using ensembleaveraged FRET (Figure S5). As expected, the apparent average FRET efficiency appeared lower than for SM-TIRF because the fluorimeter spectra included all proteins, including mislabeled molecules with only single dye labels, while the SM approach excluded mislabeled proteins using the alternating laser excitation approach (see above).42 However, the bulk FRET experiments were qualitatively consistent with SM measurements, confirming the conformational stabilization by nanobubble-laden surfaces. In particular, intact AF555CF633-NfsB exposed to a nanobubble-coated surface and a TMS-coated surface without nanobubbles exhibited FRET efficiencies of 9.1% and 5.2%, respectively (Figure 5, Table S2). Moreover, denatured AF555-CF633-NfsB exposed to a nanobubble-coated surface and a TMS-coated surface without nanobubbles exhibited FRET efficiencies of 4.5% and 4.3%, respectively. Thus, the trend in the fluorometer results further validated the SM FRET studies, demonstrating that surfacebound nanobubbles acted to preserve protein conformation in the presence of hydrophobized glass surfaces. One of the most striking results of this study is that proteins desorbing from nanobubble-covered glass are more folded than those desorbing from both methylated and unfunctionalized glass. This observation may be explained by considering the role of surface heterogeneity in protein unfolding. Weltz et al. showed that anomalous surface sites, or “hot spots,” were responsible for the majority of surface-mediated protein unfolding events.12 Similarly, nanobubble stability on hard surfaces is thought to be due to contact line pinning on structural heterogeneities on the surface. Thus, it is likely that the nanobubbles themselves attach to these same denaturing sites passivating them against subsequent protein interactions. Furthermore, the FRAP studies described above suggested that a small amount of protein adsorbed from solution to form a sacrificial layer on the nanobubble surfaces. This adlayer presumably formed a protective layer of intact protein at the nanobubble−water interface, thereby protecting subsequently adsorbing protein molecules from interactions with the gas phase. At this point, these experiments cannot determine what part of the protein sticks to the bubble; previous molecular dynamics studies have pointed to SH3 domains or hydrophobic active sites, the latter of which is more likely for an enzyme like nitroreductase.50,51 Regardless, because these denaturing sites are covered by nanobubbles and the protein adlayer, the dissolved protein molecules are blocked from interacting with these same anomalous denaturing sites, leading to a higher folded protein fraction in solution. As a result, nanobubbles paradoxically appear to protect proteins from surface mediated denaturation because of, rather than in spite of, the strong adsorption of proteins to the air−water interface.

Letter

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpclett.9b00806.

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Detailed description of materials and methods, additional figures showing flow setup of TIRF imaging, images showing photobleaching of dye-labeled NfsB, circular dichroism spectra of thermally denatured NfsB, schematic of TIRF fluorometer setup, and tables showing actual nanobubble counts and surface FRET as measured by fluorometer (PDF)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Daniel F. Kienle: 0000-0002-0962-4608 Andres F. Chaparro Sosa: 0000-0003-4094-6194 Jennifer N. Cha: 0000-0002-2840-1653 Joel L. Kaar: 0000-0002-0794-3955 Andrew P. Goodwin: 0000-0002-7284-4005 Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank the National Institute of Biomedical Imaging and Bioengineering for support of this research through grants DP2EB020401 and R21EB026006. D.F.K., J.L.K., and D.K.S. acknowledge support from the US Defense Threat Reduction Agency under award number: HDTRA1-161-0045. The authors also thank Prof. Christopher Bowman for use of his irradiation setup for creating the flow chambers.

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ABBREVIATIONS NfsB, nitroreductase; FRET, Förster resonance energy transfer; SM, single molecule; TIRF, total internal reflection fluorescence microscopy; AF488, AlexaFluor 488; AF555, AlexaFluor 555; HMDS, hexamethyldisilazane; TMS, trimethylsilyl; FRAP, fluorescence recovery after photobleaching

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REFERENCES

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