Imaging Active Surface Processes in Barnacle Adhesive Interfaces

Dec 17, 2015 - Juvenile and adult barnacles do not usually relocate. ... (b) Top-view of chamber housing a reattached barnacle on a SPRI gold-coated c...
0 downloads 0 Views 2MB Size
Download PDF
Article pubs.acs.org/Langmuir

Imaging Active Surface Processes in Barnacle Adhesive Interfaces Joel P. Golden,† Daniel K. Burden,‡ Kenan P. Fears,§ Daniel E. Barlow,§ Christopher R. So,‡ Justin Burns,‡ Benjamin Miltenberg,∥ Beatriz Orihuela,⊥ Daniel Rittshof,⊥ Christopher M. Spillmann,† Kathryn J. Wahl,*,§ and Leonard M. Tender*,† †

Center for Bio/Molecular Science and Engineering, ‡National Research Council Post Doc, §Chemistry Division, Naval Research Laboratory, Washington, D.C. 20375, United States ∥ American Society for Engineering Education, NREIP, Washington, D.C. 20036, United States ⊥ Duke University Marine Lab, Beaufort, North Carolina 28516, United States S Supporting Information *

ABSTRACT: Surface plasmon resonance imaging (SPRI) and voltammetry were used simultaneously to monitor Amphibalanus (=Balanus) amphitrite barnacles reattached and grown on gold-coated glass slides in artificial seawater. Upon reattachment, SPRI revealed rapid surface adsorption of material with a higher refractive index than seawater at the barnacle/gold interface. Over longer time periods, SPRI also revealed secretory activity around the perimeter of the barnacle along the seawater/gold interface extending many millimeters beyond the barnacle and varying in shape and region with time. Ex situ experiments using attenuated total reflectance infrared (ATR-IR) spectroscopy confirmed that reattachment of barnacles was accompanied by adsorption of protein to surfaces on similar time scales as those in the SPRI experiments. Barnacles were grown through multiple molting cycles. While the initial reattachment region remained largely unchanged, SPRI revealed the formation of sets of paired concentric rings having alternately darker/lighter appearance (corresponding to lower and higher refractive indices, respectively) at the barnacle/gold interface beneath the region of new growth. Ex situ experiments coupling the SPRI imaging with optical and FTIR microscopy revealed that the paired rings coincide with molt cycles, with the brighter rings associated with regions enriched in amide moieties. The brighter rings were located just beyond orifices of cement ducts, consistent with delivery of amide-rich chemistry from the ducts. The darker rings were associated with newly expanded cuticle. In situ voltammetry using the SPRI gold substrate as the working electrode revealed presence of redox active compounds (oxidation potential approx 0.2 V vs Ag/AgCl) after barnacles were reattached on surfaces. Redox activity persisted during the reattachment period. The results reveal surface adsorption processes coupled to the complex secretory and chemical activity under barnacles as they construct their adhesive interfaces.

1. INTRODUCTION Barnacles, marine crustaceans found in all of the world’s oceans at all depths, adhere permanently to surfaces by building a complex, layered interface of protein, cuticle, and, in many species, calcite as they expand their periphery. Acorn barnacles are protected by a hard, volcano-like shell encasing their soft body tissue. Barnacles grow by undergoing periodic molts as often as every few days.1 However, unlike other Arthropoda including insects, spiders, shrimp, crabs, and lobsters, acorn barnacles shed the cuticle on their exposed soft parts, but they do not resorb or shed their protective shell when they molt. The shell is segmented into numerous plates, and growth is at the edges of the plates. The outer shell plates are enlarged at the seams, and the shell as a whole expands in height by calcification at the bottom edges abutting the base.2−4 The process of adhesive interface development proceeds hidden under the barnacle and is intimately linked with molting.5 So while barnacles secrete adhesive “under water”, the deposition and curing processes are covered and protected by cuticular © 2015 American Chemical Society

tissue and, in many acorn barnacles, newly formed calcite in the base plate.6,7 Nonetheless, the interfaces are very hydrated, with the near-surface region having as much as 25−50% water by weight,8 and are even more hydrated (∼90% by weight)9 if the adhesive becomes very thick due to inadequate sealing or other adherence-related problems.5,10,11 The details of how the proteins in the barnacle’s buried interface are cured into highly insoluble, but hydrated, material is incomplete.12−14 Despite the significant hydration levels in the interface, barnacles are one of the most tenacious organisms in the marine environment.15 The geometry and mechanics of the calcite base plate16 may play a significant role in the adhesion,17 but even barnacles with no calcification of the base plate adhere strongly.15 On the microscopic scale, barnacle adhesive is composed of a nanofibrillar protein matrix18−20 with spectroReceived: September 1, 2015 Revised: December 13, 2015 Published: December 17, 2015 541

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

Figure 1. (a) Cross-section view of barnacle in chamber on SPRI chip showing SPRI light path (gray lines) and SPRI angle (θ). SPRI chip was mounted on prism with index matching fluid. Working (WE), counter (CE), and reference (RE) electrodes were connected to a potentiostat for electrochemical measurements. WE connection was made outside the chamber. (b) Top-view of chamber housing a reattached barnacle on a SPRI gold-coated chip. A black O-ring (2 cm diameter) forms a fluid seal between the chamber and chip.

scopic signatures showing significant β-sheet composition.19,20 The process of interface development involves multiple steps, with temporally and spatially resolved secretory activities under the base plate. Temporally resolved chemical changes contribute to adhesion during reattachment of barnacles; enhanced adhesion was correlated to changes in chemistry including increased β-sheet conformation of the proteins and increased phenolic character within the interface.6 The source and specific chemistry of these components14,21 as well as their curing mechanism22 remain incompletely understood. Secretions from the capillary ducts and cuticle during molting both contribute to the interface chemistry and curing. Here, we use surface plasmon resonance imaging (SPRI) to reveal the surface-related processes occurring during reattachment and growth of barnacles in situ. SPR (surface plasmon resonance) is highly sensitive to changes in interfacial refractive index due to deposition of substances (e.g., proteins).23 In contrast, SPR is insensitive to proteins in solution less than 300 nm away from the surface. This feature enables limits of detection on the order of 500 pg adsorbate per square centimeter.24−28 SPRI is a variation of SPR that enables optical mapping of processes at interfaces.29−31 SPRI has been used to explore temporary adhesion of settlement stage barnacles (cyprids), revealing surface exploratory behavior and deposition of protein footprints on test surfaces of varying surface chemistry.32−34 Here, SPRI reveals the different stages of barnacle reattachment: displacement of water with surface adsorbed species, followed by normal growth and development that expand the interface with additional complex surface phenomena. Both reattachment and growth are accompanied by secretions outside the barnacle. In situ cyclic voltammetry was used to evaluate the interfacial redox activity.35 Optical microscopy, scanning probe microscopy, and infrared spectroscopies were used to identify chemical features of the materials involved in interface building.

Corning, Midland, MI) coated glass panels at Duke University Marine Laboratory and reared as described by Holm et al.38 Approximately 4− 5 weeks after settlement, when barnacles were large enough to eat brine shrimp nauplii, the panels were shipped overnight to the Naval Research Laboratory and kept in 32 ppt artificial seawater (ASW, Instant Ocean, Specific Gravity 1.022) in an incubator maintained at 23 °C on a 12 h light/dark cycle. Barnacles in the incubator and in the SPRI chamber were fed brine shrimp (Artemia spp.) nauplii every other day, and the seawater was changed twice a week. Barnacles used in experiments reached maturity and had well-formed calcified base plates, but they were young enough to be vigorously growing and molting every 3−5 days. Barnacles to be used for SPRI and ATR-FTIR experiments (between 4 and 7 mm in diameter) were gently dislodged from the silicone-coated glass panels, rinsed with distilled water, and placed onto the substrates for those experiments as described below. No weights or other measures were used to hold down the barnacles 2.2. In Situ Analysis of Barnacle Interfaces. 2.2.1. SPRI. Gold surfaces were standard SPRI substrates (GWC Technologies) consisting of SF10 glass slides (25 × 38 × 1 mm3) onto which a 7 nm thick titanium adhesion layer was deposited followed by a 38 nm gold layer. Prior to use, SPRI substrates were rinsed vigorously with ethanol followed by DI water and air-dried. Electrical connections for voltammetry were made directly to the gold surface near one edge using a low-temperature soldering iron and indium solder to minimize heat damage to the gold layer. The connection was coated with 5 min epoxy (Devcon) for strain relief. A commercial surface plasmon resonance imager (SPRimager Horizon, GWC Technologies: Kretschmann configuration) with a CCD detector, operated at a manually adjustable fixed angle29 at 800 ± 6 nm, was equipped with a custom-designed and fabricated chamber to house live barnacles for experiments over days to weeks. The chamber (Figure 1) was fabricated out of poly(methyl methacrylate) (PMMA) and had a volume of 120 mL; this enabled in situ monitoring of barnacles in ASW with the gold surface of the SPRI substrate comprising the bottom of the chamber. The chamber was sealed to the gold surface by a 2 cm diameter nitrile rubber O-ring. Figure 1b shows the O-ring and 1.6 cm diameter opening exposing the gold surface to the ASW media. For experiments with voltammetry, the electrical connection to the gold surface was outside the chamber. The top of the chamber was open to the laboratory environment. The bottom of the SPRI substrate (with the attached chamber) was mounted onto the prism of the SPRI imager using index matching fluid (Cargille Master Calibration Liquid, no. 19268, n = 1.6304). SPRI spatially maps interfacial processes.29 Incident light reflected off the backside of the gold/barnacle and seawater interface (through the underlying glass) is imaged by a CCD detector. This generates a spatial image of the relative intensity of light reflected by the interface as determined by local variation in refractive index at the interface. In such images, brighter regions (high SPRI signal intensity) correspond to regions of the interface that have a higher refractive index than seawater, caused, for example, by displacement of water by adsorbed

2. MATERIALS AND METHODS 2.1. Barnacles. Juvenile and adult barnacles do not usually relocate. The approach we use here is consistent with that of our recent work where barnacles dislodged from a silicone substrate were reattached to a surface.6,7,36 This method is widely used to assess barnacle adhesion on test substrates, referred to as a reattachment assay, where barnacles are allowed a period of time (days to weeks) to readhere to surfaces before a push-off test is used to measure release stress.36 Mass cultured laboratory barnacle cyprids (Amphibalanus amphitrite) were set37 on 7.6 × 15.2 × 0.64 cm3 T2 silicone (Dow 542

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

Figure 2. Sequence of SPRI images for a barnacle on gold-coated substrate at (a) 0.2, (b) 5.0, and (c) 21 h after reattachment. Images represent the change in refractive index at the substrate interface and show interaction with the underside of the barnacle. Diameter of barnacle was ∼7 mm. Edges of the round chamber housing barnacle are present in lower section of each image. Images are distorted due to SPR imaging angle. (d) Initial growth measured by SPRI. Five regions in (c) were selected to indicate growth as measured by SPRI pixel values. Region 1: control region on gold surface (gray solid); region 2: inside barnacle edge (black solid); region 3: barnacle middle (black dashed); region 4: “aura” outside barnacle (black dotted); and region 5: control region outside the chamber indicates system drift (gray dotted). Video compilation of images showing initial reattachment is provided in Supporting Information (Video S1). protein or other higher index material. In contrast, dark regions correspond to regions that are of relatively low refractive index, caused, for example, by regions of the gold surface reduced in adsorbed substances. Other factors, including temperature and salt content39 as well as electrochemical current,40 can induce changes in the interfacial refractive index. SPR reflectivity curves have a linear region that provides positive linear correlation between refractive index and pixel intensity. To maintain pixel intensity in the linear range, the following protocols were used to set the SPR angle before each experiment. With media in the chamber while observing pixel intensity on a live image, the incident angle was set to obtain the lowest pixel intensity and then adjusted to a lower angle to increase pixel intensity by approximately 1 /4 of the maximum value minus the lowest pixel intensity. Positive linear correlation of refractive index with pixel intensity was verified using calibrated concentrations of ethanol. The linear range for this instrument is specified by the manufacturer for pixel intensity values between 75 and 185 (maximum range of 0−255, 8 bits). When the intensity exceeds 180, the correlation is still positive, but linearity is lost, yielding smaller increases in pixel intensity with increased index of refraction. Because experiments were in some cases carried out over multiple days to weeks, the angle was reset as needed to maintain response linearity. Sequential SPRI images were recorded over time to provide spatiotemporal documentation of barnacle growth from the perspective of changes in refractive index of the gold/barnacle and seawater interface. Images were captured at 1 s intervals during early stages of reattachment and at 5 min intervals thereafter. The incident angle of light determined the range of intensities covered by the dynamic range of the SPR image. Since intensity of reflected light by the barnacle/gold interface changed dramatically over time, not all features of the interface could be imaged using the same incident angle without saturating the CCD detector. As a consequence, some features described here were captured using different incident angles during the

SPRI observations. The incident angle was set at the beginning of each experiment to a fixed value that would best capture the intended highlighted features, as described above. The SPRI images have an elongated width due to the CCD detector observation angle; fiducial marks on the substrate were used to compare in situ SPR images with ex situ microscopy. Nominal pixel resolution of the SPRI is 16 × 32 μm2 (vertical × horizontal), determined by resolution of the CCD camera. 2.2.2. Voltammetry. The SPRI chamber was equipped with an electrochemical reference electrode (Ag/AgCl, 3 M KCl; Bioanalytical Systems, Inc.) and a counter electrode (titanium bolt, United Titanium), enabling simultaneous electrochemical experimentation using the SPRI substrate as the working electrode. In all experiments reported here, the working electrode was at open circuit, i.e., no potential was applied while the barnacle was growing except when performing cyclic voltammetry (CV). CV was performed every 12 h whereby the potential of the gold substrate was changed from −0.3 to 0.4 V and back to −0.3 V (vs Ag/AgCl) at 10 mV/s twice. All voltammetric experiments were performed with a software controlled (Multistat, Scribner Assoc.) potentiostat (1470E, AMETEK). Similar to the SPR imaging, thermal and salinity changes can impact electrochemical current.40 2.2.3. ATR-FTIR Spectroscopy. A 10 × 50 × 2 mm3 Ge ATR prism in a liquid-cell holder was used to monitor surface chemical processes of reattached barnacles in ASW. ATR-FTIR spectra were obtained using a Thermo-Fisher 6700 spectrometer coupled with a Harrick Horizon ATR accessory and equipped with a liquid nitrogen cooled mercury−cadmium−telluride (MCT) detector. Germanium prisms were cleaned by sonication in detergent and DI water. The liquid cell holder was filled with ASW, and background spectra were collected. Barnacles were removed from silicone panels and immediately placed in contact with the germanium ATR prism similar to that for the SPRI experiments, except that six barnacles were attached simultaneously to cover a large fraction of the long ATR prism. Spectra were obtained 543

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

Figure 3. (a) ATR-FTIR spectrum ∼5 min after barnacle reattachment on the Ge ATR prism in ASW. The spectrum indicates displacement of water and adsorption of protein at the surface. (b) ATR-FTIR protein adsorption profile over ∼170 h (∼7 days) following barnacle reattachment onto Ge in ASW. The amide II absorbance at 1545 cm−1, normalized by total barnacle area at the surface, was used as a relative measure of protein adsorption. and referenced to ASW. Protein adsorption profiles were obtained by plotting the amide II absorption intensity (1545 cm−1), normalized for total barnacle area at the surface, versus time. 2.3. Ex Situ Analysis of Barnacle Interfaces. Separate analyses were performed after the in situ studies to identify the band features observed by SPRI. These included optical microscopy, atomic force microscopy (AFM), and imaging FTIR in both reflectance (barnacles grown on gold-coated SPR chips) and transmission (barnacles grown on CaF2) modes. 2.3.1. Optical Microscopy. Following SPRI experiments, goldcoated SPRI substrates with adhered barnacles were prepared for optical microscopy by removing the internal soft barnacle bodies and immersing the attached shells in 0.1 M ethylenediaminetetraacetic acid (EDTA, Sigma-Aldrich) solution to demineralize barnacle side and base plates. The demineralization process took up to 72 h with the immersion solution being replaced with fresh EDTA solution daily. Once fully demineralized, all organic matter above the cuticular layer was carefully removed and the remaining barnacle plaques were imaged in bright-field reflection mode using a Nikon AZ100 microscope to correlate features from the two imaging techniques. 2.3.2. Atomic Force Microscopy. Additional barnacle base plate regions were examined by AFM after demineralizing with EDTA to remove the calcareous base plate. This experiment was performed for barnacles grown on CaF2 substrates to facilitate transmission spectroscopy experiments. In some regions, the cuticle remaining after etching was peeled back from the substrate to reveal the underlying chemistry. AFM images were recorded in intermittent contact mode (Veeco Nanoman, Bruker Nanoscope V controller). Uncoated n-doped silicon tips (TESP, Bruker) with a 42 N/m nominal spring constant and typical apex radius of 8 nm were used at ca. 350 kHz resonance frequency. Images were captured at a resolution of 512 × 256 at a rate of 0.6 Hz. 2.3.3. Imaging FTIR Spectroscopy. Demineralized barnacle bases on gold-coated SPRI substrates were analyzed with a vertex 70v spectrometer (Bruker) equipped with a Hyperion microscope. Optical and FTIR images were collected at 15× using a focal plane array IR detector. IR spectra were acquired in reflectance geometry at 8 cm−1 resolution for 1024 scans, using a clean gold-coated SPRI substrate as the background. Regions of the demineralized bases examined by AFM as described above were also analyzed with a 6700 FTIR spectrometer (Thermo Scientific, Waltham MA) outfitted with a continuum microscope and MCT detector. Transmission spectra were acquired at 4 cm−1 resolution for 256 scans. The aperture size was set at 20 × 20 μm2, and a cleaned area of the CaF2 substrate outside the barnacle periphery served as the background.

were investigated, and each of the features described below was observed for at least 3, but typically many more, barnacles. All barnacles were 4−7 mm diameter at reattachment. Figure 2a−c depicts a time sequence of SPRI images acquired immediately after reattachment of a barnacle onto an SPRI substrate but prior to additional measurable growth (21 h). A clear increase (brightening) in SPRI signal intensity is observed for the area underneath the barnacle basis. The overall increase is uniform, with average signal increasing monotonically over time (Figure 2d). A majority of the surface brightening was observed directly under the barnacle, with additional increase in intensity at the barnacle periphery. The intensity of the region outside the chamber did not change with time, whereas the intensity of the seawater region outside the barnacle base plate area increased gradually over time, having a lower intensity and slope than the regions associated directly with the barnacle and sometimes being quite bright at the periphery. The SPRI technique is highly sensitive to changes in interfacial refractive index due to proximity and/or deposition of substances at interfaces. The initial placement of the barnacle on the SPRI chip resulted in only a modest change in intensity; in contrast, with time the intensity changes under the barnacle increased substantially over the background signal. This suggests that during barnacle reattachment the seawater under the barnacle is gradually replaced over a period of hours with material from the barnacle having a higher refractive index than that of seawater. The process is not always uniform, but it typically proceeded from the periphery inward, eventually filling in the entire area under the barnacle (see Supporting Information Video S1, showing the intensity changes during reattachment). We performed a similar experiment by reattaching barnacles on a germanium prism exposed to seawater and performing time-resolved ATR-FTIR to monitor the chemical changes at the surface. The ASW background spectrum obtained before barnacles were placed on the prism showed bands expected for water.8 The absorbance spectra after barnacle attachment revealed protein adsorption and water displacement within 5 min (Figure 3a). There was a decrease in the 3000−3500 cm−1 region (OH, blue shaded region), as well as increases in spectral regions associated with protein amide I, II, and III regions (pink shaded regions between 1200 and 1700 cm−1). The shape of the OH region also indicated amide (NH) contributions were increasing (pink shaded area ∼3300 cm−1). The spectra are consistent with previous studies of barnacle adhesive interface

3. RESULTS AND DISCUSSION 3.1. Initial Barnacle Reattachment. Barnacle viability was confirmed by observing barnacle growth (expanding basis), feeding, and molting after reattachment. A total of 41 barnacles 544

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

Figure 4. Sequence of SPRI images highlighting barnacle growth and formation of concentric ring-like pattern in the new growth region beneath barnacle at (a) 3 h, (b) 10 days, and (c) 14 days after the barnacle was first reattached on the substrate. Beginning diameter of barnacle was ∼6 mm. Video compilation of images showing growth is provided in Supporting Information (Video S2).

Figure 5. Comparison of SPRI and optical microscopy images: (a) SPRI image, (b) optical micrograph of the same barnacle after demineralization, (c) the two images shown side by side, and (d) close-up of the inset in (c) showing coincidence of the SPRI bands and ecdysal lines and ducts in the microscopy image. Images were aligned using fiducial marks in the gold surface.

chemistry of undisturbed8 and reattached6 barnacles. Since the amide I band and the primary water band overlap, the amide II band intensity was used to monitor the spectral changes as a function of time over a period of several days. The results are presented in Figure 3b. While this experiment differs from the SPR experiment in that there were multiple barnacles (5) and different substrate materials (Ge vs Au coated glass), the results are consistent with the SPR studies: rapid and increasing adsorption of barnacle proteins to the surface at the edges and under the barnacle, with simultaneous displacement of water from the surface. Whether the surface adsorption process is a result of gradual relaxation of the barnacle base plate against the surface, new secretory activity, or a combination of both is not

known. In many instances, we saw evidence for secretory activity during reattachment. Regardless, chemistries consistent with proteins are quickly in intimate contact with the surface under the barnacle during reattachment. 3.2. Barnacle Growth. The imaging SPR system provided an opportunity to examine the development of barnacle interfaces after reattachment. Barnacles were monitored over a period of several weeks. During this period, barnacle base plate diameters increased by 1−2 mm, and we tracked the number of times, and when, the barnacles cast off molts. Figure 4 depicts a representative time sequence of SPRI images of growth of a reattached barnacle. Here, barnacle growth is indicated by the expanding bright region relative to the 545

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir seawater background intensity; the diameter increased at a rate of approximately 0.21 mm/day over 14 days. The reattachment region was generally very bright and often relatively uniformly bright. In contrast, new interface development was accompanied by formation of distinct patterns of bright and dark rings (Figure 4b,c) that appeared only in the regions of new growth. Some regions of the base plate show radial spoke-like regions with signal intensities similar to the bright central region. There are also persistent bright and dark features that are maintained at and move along with the growing perimeter, as well as irregular-shaped bright regions that fluctuate and persist just beyond the growing perimeter (see Supporting Information Video S2 for a video showing barnacle growth). Subsequent imaging by ex situ SPRI and optical microscopy (Figure 5a,b) of the same barnacle/substrate interface, after removal of the barnacle body and base plate calcite and tissue above the cuticle layer, revealed that the pairs of bright/dark rings coincide with ecdysal lines and capillary ducts (Figure 5c,d) formed with each molting cycle during growth (see Supporting Information Video S3 for direct comparison of the two imaging methods). Both the circumferential bands and radial spoke-like regions were brighter due to increased white light scattering in optical microscope images and were consistently correlated with the bright regions in the SPRI images. Analysis by ex situ imaging FTIR revealed that the banded structure was associated with a radial variation in chemical composition, with the brighter banding pattern strongly correlated with the presence of amide components (Figure 6). While the entire base plate region revealed an abundance of amide chemistries, the banded regions with highest amide intensity were found at radial locations just beyond the capillary duct orifices. On a separate barnacle reattached to CaF2, we carefully peeled back some of the cuticle layer to reveal the continuous radial bands sandwiched between the cuticle and the substrate (Figure 7a). The material remained adhered to the substrate and was spectroscopically consistent with protein (Figure 7b), whereas the regions of cuticle between bands were released with little remaining residue. AFM (Figure 7c,d) revealed that the regions with adhered material were composed of an interlocked fibrillar structure. Our prior spectroscopic observations both in situ8 and ex situ6,7,20 revealed that the dominant interfacial chemistry nearest the surface is proteinaceous and is often composed of nanofibrillar networks. We note that the AFM reveals the upper surface of the cohesive failure and may have some cuticle residue; however, the images clearly reveal that the cuticle layer is secured to the substrate in a chemically discontinuous manner. The banded adhesive failure pattern is intriguing in light of adhesion mechanics predictions that mechanical discontinuities and stiffness variation can impart crack-toughening properties to the interface.41−43 The banded ring pattern in Figure 5 coincides with the orifices of ducts that play an important role in barnacle adhesive deposition.6,7 Given the protein-rich band positions are just beyond the duct orifices (Figures 6 and 7), it is plausible that the protein-rich bands are secreted directly from the ducts. The time-lapse images of barnacle growth reveal more details of the development of the bright and dark ring features under barnacles. The leading edge of the barnacle periphery typically appears bright, with a similar intensity as the brighter bands. The dark bands appear from a splitting of the bright band at the growing edge, leaving a dark band behind as a portion of the bright band continues to expand radially. This can be seen in

Figure 6. FTIR spectra (a) from two regions of demineralized barnacle base plate noted in marked circular areas in (b). The map (b) shows in grayscale the absorbance intensity variation in the amide II band across a section of base plate. The lighter gray banded regions (red dot and corresponding upper spectrum) and darker regions (blue dot and corresponding lower spectrum) are clearly demarcated at radial positions associated with the terminal regions of the capillary ducts.

the set of images shown in Figure 8, where this process is shown occurring twice, between the first two and then the last two frames shown. New pairs of bands were observed to coincide with each time the barnacle molted. We hypothesize that the darker bands arise from fracture, and then expansion, of the cuticle layer during molting as the side plates expand and stretch and pull the cuticle layer below along the surface. This is an integral process to barnacle interface development whereby the newly matured cuticle, residing folded accordion-like above the prior, hardened cuticle, is exposed to the surface when the old cuticle breaks. The fluids released in this process, called ecdysal fluids, contain enzymes and other chemistries related to cuticle thinning.44,45 The band under this newly formed region may be darker if these fluids react with the cuticle rather than adsorb to the surface or if the new cuticle is not in full contact with the gold surface. The increased brightness in the area freshly exposed to molting fluids and chemistries from the ducts suggest that the materials coming out have a high affinity for the surface or higher index of refraction. While we are aware of many proteins that have been identified and associated with barnacle adhesion,21 we do not yet know specifically which proteins are localized to the bright regions or their chemical functionality and surface affinity. We do not expect to see strong effects from the calcite layer above, which is more than a micrometer away from the substrate in a live barnacle interface and not anticipated to contribute significantly to signal in the 546

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

Figure 7. (a) Optical micrograph (ca. 340 × 260 μm2) of exposed sub-baseplate band and AFM tip position with respect to the band edge. Region imaged by AFM is highlighted in red. (b) Transmission IR microscopy of band structure from (a) showing a Fourier transformed spectrum with major amide I/II components typical for proteins. (c) Intermittent contact AFM image (15 × 15 μm2) of band edge with height scale and (d) magnified region (5 × 5 μm2) showing off-band surface features to be 10−20 nm in height.

Figure 8. Sequential frames showing progression of banding pattern seen under barnacle with SPRI in 14 days of growth. The perimeter of the growing barnacle typically has a narrow dark line, whereas just inside the expanding perimeter is a lighter band. As the barnacle grows, the bright band at the far right perimeter of the barnacle can be seen to split into two bright bands separated by a darker region. The inner bright band remains in place, whereas the outer band moves laterally as the barnacle base plate expands. This process happens twice in the series of images, between frames a and b and frames c and d. Yellow lines indicate the final width of the dark rings.

In the SPRI experiments, we noticed secretory activity at the periphery outside the barnacle, with a higher index material radiating from the barnacle and extending outward over many hours. While secretory activity like this often occurred during reattachment, and might plausibly be related to wounding of the barnacle, we also observed secretory activity at the periphery for all barnacles during normal growth and development. The secretions appeared periodically, but we were not able to associate them with any particular event during base plate development and they may, therefore, be unrelated. It is not known where the secretions originate, but similar secretions have been observed previously;10 they are possibly

SPR experiments; this is consistent with our prior in situ ATRFTIR measurements.8 Interestingly, the region under the leading edge remains bright through this process, suggesting that the adsorbed secretions are carried along with the movement of cuticle attached to the side plates. We found previously that proteinaceous secretions extend all the way to the periphery of the barnacle under the leading edge of the base plate, under the newly developing cuticle.7 These secretions are in place well in advance of the formation of the outermost new capillaries and any secretory activity and may arise from the cuticle itself. The results suggest that the molting and capillary secretion processes may result in desorbing or reworking these protein layers on the surface. 547

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

form associated with the barnacle.35 Additional control CVs in which SPRI substrates were divided into two electrically separate regions with a barnacle reattached to only one region exhibited the observed redox activity for both regions, which we infer to indicate that the redox active compound may be soluble and not restricted to the barnacle/substrate interface (not shown). What processes involved in growth or adhesion might be responsible for the observed redox response? While the mechanism for barnacle adhesive curing is incompletely understood and may include the observed redox response, it is well-known that chemistries of cuticle development and hardening, called sclerotization, are also present in barnacle adhesive interfaces.5,49 We speculate that these chemistries may be responsible for the observed redox response. For example, the chemistries required for sclerotization are processed by enzymes and include precursor molecules like 3,4,-dihydroxyphenylalanine (DOPA), which are transformed by decarboxylation to dopamine and then into either N-acetyl dopamine (NADA) or N-β-alanyldopamine (NBAD).50,51 These molecules are associated with the cuticle color and perhaps mechanical properties, with NBAD associated with darker cuticle and NADA associated with lighter colors.51,52 The oxidation and reduction potentials for both these molecules fall in the same range observed here, with oxidation around 0.22 V and reduction at ∼0.1 V,53 and differ markedly from those of DOPA.54 Barnacles, and in particular the species examined in this study, A. amphitrite, typically have very transparent cuticle membranes, implying tanning chemistries may be associated with NADA. Oxidase activity has long been noted in barnacle interfaces5,55,56 but without evidence of post-translationally modified proteins. With the difficulty of monitoring chemistry in a buried interface, the questions of what molecules are crosslinked, the chemistry of the cross-links, and what relation the redox activity bears to adhesion remain open.

either adhesive proteins or related to water-borne settlementinducing proteins.46,47 Finally, we sometimes noted a dark ring around the barnacle perimeter. We do not know whether this is a region cleared by the barnacle either mechanically by the shell side plates or chemically or whether it is an optical artifact of the experiment (see Supporting Information Video S4 for a closeup of the expanding barnacle perimeter). One possibility is that these features of the advancing leading edge may be related to secretions underneath the advancing cuticle that prepares the surface for adhesion. The observation of possible preparation of the surface by the advancing perimeter proposed here is reminiscent of the recent report by Gohad et al. in which barnacle cyprids secrete both protein and lipids as part of the transition from temporary to permanent adhesion.48 However, we note that both protein and lipids would have higher index of refraction than ASW and would, therefore, not account for a dark band at the far periphery. Thus, if the observation is not an artifact, either chemical or mechanical clearing of the surface is more probable. 3.3. Cyclic Voltammetry. For some of the experiments, the electrochemical state of SPRI surfaces exposed to barnacles was monitored. Figure 9 depicts two CVs recorded at 120 (Figure 9a) and 180 h (Figure 9b) after reattachment of a barnacle. The featureless CV depicted in Figure 9a is consistent with CV of controls lacking barnacles (ASW medium with and without brine shrimp). In contrast, CV depicted in Figure 9b indicates abundance of redox active compounds in the reduced

4. CONCLUSIONS In summary, we were able to resolve several distinct stages of the reattachment and growth of barnacles on gold surfaces in artificial seawater. After reattachment, over a period of several hours to days, the barnacle develops intimate contact with the substrate, displacing water with proteinaceous secretions under the base plate. After a period of latency, the barnacle expands its base plate. Development of new base plate is associated with formation of sets of alternating dark and bright bands in the SPR image, indicating regions with varying refraction indices. The brighter regions were associated with adsorbed protein in regions near rings of adhesive capillary ducts that form with each molt cycle. The darker bands were identified with regions with significantly lower protein content in ex situ studies and were associated with the regions of newly formed cuticle that expand as the barnacle base plate develops. The leading edge of the barnacle base plate remains bright, consistent with observation of adsorbed proteins between the surface and cuticle in these regions. Secretory activity outside the barnacle was also noted, which came and went in waves and did not appear to be necessarily associated with any process occurring in the interface; these materials were adsorbed to, and mobile on, the surface and may be related to settlement cues and pheromone activity. Finally, CV was well-resolved and consistent with redox activity of known chemistries associated with cuticle tanning. Whether

Figure 9. Cyclic voltammetry of barnacles using gold SPRI substrate as working electrode. Shown are cyclic voltammograms at 120 h (a) and 180 h after reattachment (b). Dashed line is the immediate second run. (a) Performed by sweeping the potential of the gold substrate twice in a row from 0.0 to −0.3 to 0.4 V and back to 0.0 V vs Ag/AgCl at 10 mV/s. (b) Performed by sweeping the potential of the gold substrate twice in a row from −0.2 to −0.3 to 0.4 V and back to −0.2 V vs Ag/AgCl at 10 mV/s. 548

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

(9) Ramsay, D. B.; Dickinson, G. H.; Orihuela, B.; Rittschof, D.; Wahl, K. J. Base plate mechanics of the barnacle Balanus amphitrite. Biofouling 2008, 24 (2), 109−118. (10) Wendt, D. E.; Kowalke, G. L.; Kim, J.; Singer, I. L. Factors that influence elastomeric coating performance: the effect of coating thickness on basal plate morphology, growth and critical removal stress of the barnacle Balanus amphitrite. Biofouling 2006, 22 (1), 1−9. (11) Holm, E. R.; Kavanagh, C. J.; Orihuela, B.; Rittschof, D. Phenotypic variation for adhesive tenacity in the barnacle Balanus amphitrite. J. Exp. Mar. Biol. Ecol. 2009, 380, 61−67. (12) Kamino, K.; Odo, S.; Maruyama, T. Cement proteins of the acorn barnacle, Megabalanus rosa. Biol. Bull. 1996, 190 (3), 403−409. (13) Naldrett, M. J.; Kaplan, D. L. Characterization of barnacle (Balanus eburneus and B. cenatus) adhesive proteins. Mar. Biol. 1997, 127 (4), 629−635. (14) Kamino, K. Mini-review: Barnacle adhesives and adhesion. Biofouling 2013, 29 (6), 735−749. (15) Yule, A. B.; Walker, G. Adhesion in Barnacles. In Barnacle Biology; Southward, A. J., Ed.; A.A. Balkema: Rotterdam, 1987; pp 389−402. (16) Hui, C. Y.; Long, R.; Wahl, K. J.; Everett, R. K. Barnacles resist removal by crack trapping. J. R. Soc., Interface 2011, 8 (59), 868−879. (17) Chung, J. Y.; Chaudhury, M. K. Soft and hard adhesion. J. Adhes. 2005, 81 (10−11), 1119−1145. (18) Dickinson, G. H.; Vega, I. E.; Wahl, K. J.; Orihuela, B.; Beyley, V.; Rodriguez, E. N.; Everett, R. K.; Bonaventura, J.; Rittschof, D. Barnacle cement: a polymerization model based on evolutionary concepts. J. Exp. Biol. 2009, 212 (21), 3499−3510. (19) Sullan, R. M. A.; Gunari, N.; Tanur, A. E.; Chan, Y.; Dickinson, G. H.; Orihuela, B.; Rittschof, D.; Walker, G. C. Nanoscale structures and mechanics of barnacle cement. Biofouling 2009, 25 (3), 263−275. (20) Barlow, D. E.; Dickinson, G. H.; Orihuela, B.; Kulp, J. L.; Rittschof, D.; Wahl, K. J. Characterization of the Adhesive Plaque of the Barnacle Balanus amphitrite: Amyloid-Like Nanofibrils Are a Major Component. Langmuir 2010, 26 (9), 6549−6556. (21) Kamino, K. Barnacle underwater attachment. In Biological Adhesives; Smith, A. M., Callow, J. A., Eds.; Springer: Berlin, Germany, 2006; pp 145−166. (22) Power, A. M.; Klepal, W.; Zheden, V.; Jonker, J.; McEvilly, P.; Byern, J. Mechanisms of adhesion in adult barnacles. In Biological Adhesive Systems; Byern, J., Grunwald, I., Eds.; Springer: Vienna, 2010; pp 153−168. (23) Homola, J.; Yee, S. S.; Gauglitz, G. Surface plasmon resonance sensors: review. Sens. Actuators, B 1999, 54 (1−2), 3−15. (24) Kunz, U.; Katerkamp, A.; Renneberg, R.; Spener, F.; Cammann, K. Sensing fatty acid binding protein with planar and fiber-optical surface plasmon resonance spectroscopy devices. Sens. Actuators, B 1996, 32 (2), 149−155. (25) Haussling, L.; Ringsdorf, H.; Schmitt, F. J.; Knoll, W. Biotinfunctionalized self-assembled monolayers on gold - surface plasmon optical studies of specific recognition reactions. Langmuir 1991, 7 (9), 1837−1840. (26) Sigal, G. B.; Mrksich, M.; Whitesides, G. M. Using surface plasmon resonance spectroscopy to measure the association of detergents with self-assembled monolayers of hexadecanethiolate on gold. Langmuir 1997, 13 (10), 2749−2755. (27) Schwarz, T.; Yeung, D.; Hawkins, E.; Heaney, P.; McDougall, A. Detection of nucleic-acid hybridization using surface-plasmon resonance. Trends Biotechnol. 1991, 9 (1), 339−340. (28) Peterlinz, K. A.; Georgiadis, R. In situ kinetics of self-assembly by surface plasmon resonance spectroscopy. Langmuir 1996, 12 (20), 4731−4740. (29) Nelson, B. P.; Grimsrud, T. E.; Liles, M. R.; Goodman, R. M.; Corn, R. M. Surface plasmon resonance imaging measurements of DNA and RNA hybridization adsorption onto DNA microarrays. Anal. Chem. 2001, 73 (1), 1−7. (30) Wang, W.; Wang, S. P.; Liu, Q.; Wu, J.; Tao, N. J. Mapping Single-Cell-Substrate Interactions by Surface Plasmon Resonance Microscopy. Langmuir 2012, 28 (37), 13373−13379.

this activity is exclusive to cuticle tanning or directly related to barnacle cement curing is a question for further investigation.

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.5b03286. Reattachment: time sequence of SPRI images for a barnacle on gold SPRI substrate up to 21 h after reattachment(Video S1; AVI) Further growth: time sequence of SPRI images showing growth and formation of rings up to 14 days after reattachment (Video S2; AVI) Comparison of SPRI and optical microscopy images: overlay transparency of SPRI image is adjusted between 0 and 100% to highlight the coincident patterns (Video S3; AVI) Barnacle leading edge during growth: sequence of highcontrast SPRI images zoomed in on the growing edge, with the three different examples showing time lapse at approximately 7 h/s (Video S4; AVI)

■

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (K.J.W.). *E-mail: [email protected] (L.M.T.). Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We acknowledge support from the Office of Naval Research Coatings Program (NRL N0001414WX20573 and N0001414WX00736; Duke N0014-11-1-0180 and N0001412-1-0365), and NRL Base 6.1 Funding. D.K.B. and J.B. were National Research Council Postdoctoral Associates.

■

REFERENCES

(1) Costlow, J. D.; Bookhout, C. G. Molting and shell growth in Balanus amphitrite niveus. Biol. Bull. 1956, 110 (2), 107−116. (2) Bourget, E.; Crisp, D. J. An Analysis of the Growth Bands and Ridges of Barnacle Shell Plates. J. Mar. Biol. Assoc. U. K. 1975, 55, 439−461. (3) Bocquet-Védrine, J. Étude du tégument de la mue chez le cirripède opercule Elminius modestus Darwin. Archives de Zoologie Expérimentale et Générale 1965, 105, 30−76. (4) Bourget, E. Barnacle shells: composition, structure, and growth. In Barnacle Biology; Southward, A. J., Ed.; A.A. Balkema: Rotterdam, 1987; Vol. 5, pp 267−285. (5) Saroyan, J. R.; Lindner, E.; Dooley, C. A. Repair and Reattachment in the Barnacle as Related to their Cementing Mechanism. Biol. Bull. 1970, 139, 333−350. (6) Burden, D. K.; Barlow, D. E.; Spillmann, C. M.; Orihuela, B.; Rittschof, D.; Everett, R. K.; Wahl, K. J. Barnacle Balanus amphitrite Adheres by a Stepwise Cementing Process. Langmuir 2012, 28 (37), 13364−13372. (7) Burden, D. K.; Spillmann, C. M.; Everett, R. K.; Barlow, D. E.; Orihuela, B.; Deschamps, J. R.; Fears, K. P.; Rittschof, D.; Wahl, K. J. Growth and development of the barnacle Amphibalanus amphitrite: time and spatially resolved structure and chemistry of the base plate. Biofouling 2014, 30 (7), 799−812. (8) Barlow, D. E.; Dickinson, G. H.; Orihuela, B.; Rittschof, D.; Wahl, K. J. In situ ATR-FTIR characterization of primary cement interfaces of the barnacle Balanus amphitrite. Biofouling 2009, 25 (4), 359−366. 549

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550

Article

Langmuir

reactions of their quinones with N-acetylcysteine and thiourea. Arch. Biochem. Biophys. 1998, 352 (1), 19−30. (54) Gorman, M. J.; Sullivan, L. I.; Nguyen, T. D. T.; Dai, H.; Arakane, Y.; Dittmer, N. T.; Syed, L. U.; Li, J.; Hua, D. H.; Kanost, M. R. Kinetic properties of alternatively spliced isoforms of laccase-2 from Tribolium castaneum and Anopheles gambiae. Insect Biochem. Mol. Biol. 2012, 42 (3), 193−202. (55) Lindner, E.; Dooley, C. A. Chemical bonding in cirriped adhesive. Proceedings of the Third International Congress an Marine Corrosion and Fouling, Gaithersburg, MD, October 2−6, 1972; pp 653−673, DOI: 10.1002/iroh.19790640304. (56) Dickinson, G. H. Barnacle Cement: A Polymerization Model Based on Evolutionary Concepts. Ph.D. Thesis, Duke University, 2008

(31) Abadian, P. N.; Goluch, E. D. Surface plasmon resonance imaging (SPRi) for multiplexed evaluation of bacterial adhesion onto surface coatings. Anal. Methods 2015, 7 (1), 115−122. (32) Andersson, O.; Ekblad, T.; Aldred, N.; Clare, A. S.; Liedberg, B. Novel application of imaging surface plasmon resonance for in situ studies of the surface exploration of marine organisms. Biointerphases 2009, 4 (4), 65−68. (33) Aldred, N.; Ekblad, T.; Andersson, O.; Liedberg, B.; Clare, A. S. Real-Time Quantification of Microscale Bioadhesion Events In situ Using Imaging Surface Plasmon Resonance (iSPR). ACS Appl. Mater. Interfaces 2011, 3 (6), 2085−2091. (34) Maleshlijski, S.; Sendra, G. H.; Aldred, N.; Clare, A. S.; Liedberg, B.; Grunze, M.; Ederth, T.; Rosenhahn, A. Imaging SPR combined with stereoscopic 3D tracking to study barnacle cyprid−surface interactions. Surf. Sci. 2016, 643, 172−177. (35) Bard, A. J.; Faulkner, L. R. Electrochemical Methods: Fundamentals and Applications, 2nd ed.; John Wiley & Sons, Inc: New York, 2001. (36) Rittschof, D.; Orihuela, B.; Stafslien, S.; Daniels, J.; Christianson, D.; Chisholm, B.; Holm, E. Barnacle reattachment: a tool for studying barnacle adhesion. Biofouling 2008, 24 (1), 1−9. (37) Rittschof, D.; Clare, A. S.; Gerhart, D. J.; Mary, A.; Bonaventura, J. Barnacle in vitro Assays for Biologically Active Substances: Toxicity and Settlement Inhibition Assays Using Mass Cultured Balanus amphitrite amphitrite Darwin. Biofouling 1992, 6 (2), 115−122. (38) Holm, E. R.; Orihuela, B.; Kavanagh, C. J.; Rittschof, D. Variation among families for characteristics of the adhesive plaque in the barnacle Balanus amphitrite. Biofouling 2005, 21 (2), 121−126. (39) Quan, X. H.; Fry, E. S. Empirical-equation for the index of refraction of seawater. Appl. Opt. 1995, 34 (18), 3477−3480. (40) Shan, X. N.; Patel, U.; Wang, S. P.; Iglesias, R.; Tao, N. J. Imaging Local Electrochemical Current via Surface Plasmon Resonance. Science 2010, 327 (5971), 1363−1366. (41) Kendall, K. Control of Cracks by Interfaces in Composites. Proc. R. Soc. London, Ser. A 1975, 341 (1627), 409−428. (42) Chung, J. Y.; Chaudhury, M. K. Roles of discontinuities in bioinspired adhesive pads. J. R. Soc., Interface 2005, 2, 55−61. (43) Xia, S.; Ponson, L.; Ravichandran, G.; Bhattacharya, K. Toughening and Asymmetry in Peeling of Heterogeneous Adhesives. Phys. Rev. Lett. 2012, 108 (19), 196101. (44) Klowden, M. J. Physiological Systems in Insects, 3rd ed.; Academic Press: San Diego, CA, 2013. (45) Gilbert, L. I. Insect Molecular Biology and Biochemistry, 1st ed.; Elsevier: Waltham, MA, 2012. (46) Clare, A. S.; Matsumura, K. Nature and perception of barnacle settlement pheromones. Biofouling 2000, 15 (1−3), 57−71. (47) Dreanno, C.; Kirby, R. R.; Clare, A. S. Smelly feet are not always a bad thing: the relationship between cyprid footprint protein and the barnacle settlement pheromone. Biol. Lett. 2006, 2 (3), 423−425. (48) Gohad, N. V.; Aldred, N.; Hartshorn, C. M.; Lee, Y. J.; Cicerone, M. T.; Orihuela, B.; Clare, A. S.; Rittschof, D.; Mount, A. S. Synergistic roles for lipids and proteins in the permanent adhesive of barnacle larvae. Nat. Commun. 2014, 5, 5−4414. (49) Lindner, E. The attachment of macrofouling invertebrates. In Marine Biodeterioration: An Interdisciplinary Study; Costlow, J. D., Tipper, R. C., Eds.; US Naval Institute: Annapolis, MD, 1984; pp 183−201. (50) Andersen, S. O. Insect cuticular sclerotization: A review. Insect Biochem. Mol. Biol. 2010, 40 (3), 166−178. (51) Andersen, S. O. Cuticular sclerotization and tanning. In Insect Molecular Biology and Biochemistry, 1st ed.; Gilbert, L. I., Ed.; Academic Press: Waltham, MA, 2012; pp 167−192. (52) Koch, P. B.; Behnecke, B.; Weigmann-Lenz, M.; FrenchConstant, R. H. Insect pigmentation: Activities of beta-alanyldopamine synthase in wing color patterns of wild-type and melanic mutant swallowtail butterfly Papilio glaucus. Pigm. Cell Res. 2000, 13, 54−58. (53) Huang, X.; Xu, R. D.; Hawley, M. D.; Hopkins, T. L.; Kramer, K. J. Electrochemical oxidation of N-acyldopamines and regioselective 550

DOI: 10.1021/acs.langmuir.5b03286 Langmuir 2016, 32, 541−550