Barnacle Balanus amphitrite Adheres by a Stepwise Cementing

Jun 21, 2012 - *E-mail: [email protected], phone: (202) 767-5419., † ... The short time window for BCS2 secretion relative to the overall ar...
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Barnacle Balanus amphitrite Adheres by a Stepwise Cementing Process Daniel K. Burden,†,‡ Daniel E. Barlow,‡ Christopher M. Spillmann,§ Beatriz Orihuela,∥ Daniel Rittschof,∥ R. K. Everett,⊥ and Kathryn J. Wahl*,‡ †

National Research Council postdoc, ‡Chemistry Division, Code 6176, §Center for Biomolecular Science and Engineering, Code 6930, and ⊥Materials Science and Technology Division, Code 6300, U.S. Naval Research Laboratory, Washington, DC 20375-5342, United States ∥ Duke University Marine Laboratory, Beaufort, North Carolina 28516, United States S Supporting Information *

ABSTRACT: Barnacles adhere permanently to surfaces by secreting and curing a thin interfacial adhesive underwater. Here, we show that the acorn barnacle Balanus amphitrite adheres by a two-step fluid secretion process, both contributing to adhesion. We found that, as barnacles grow, the first barnacle cement secretion (BCS1) is released at the periphery of the expanding base plate. Subsequently, a second, autofluorescent fluid (BCS2) is released. We show that secretion of BCS2 into the interface results, on average, in a 2-fold increase in adhesive strength over adhesion by BCS1 alone. The two secretions are distinguishable both spatially and temporally, and differ in morphology, protein conformation, and chemical functionality. The short time window for BCS2 secretion relative to the overall area increase demonstrates that it has a disproportionate, surprisingly powerful, impact on adhesion. The dramatic change in adhesion occurs without measurable changes in interface thickness and total protein content. A fracture mechanics analysis suggests the interfacial material’s modulus or work of adhesion, or both, were substantially increased after BCS2 secretion. Addition of BCS2 into the interface generates highly networked amyloid-like fibrils and enhanced phenolic content. Both intertwined fibers and phenolic chemistries may contribute to mechanical stability of the interface through physically or chemically anchoring interface proteins to the substrate and intermolecular interactions. Our experiments point to the need to reexamine the role of phenolic components in barnacle adhesion, long discounted despite their prevalence in structural membranes of arthropods and crustaceans, as they may contribute to chemical processes that strengthen adhesion through intermolecular cross-linking.

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Like mussels and sandcastle worms, barnacles adhere to surfaces with protein-rich cement they secrete and cure underwater.8−13 Despite considerable taxonomic and histological observations dating to Darwin and his contemporaries,14,15 the complex physiological processes involved in creating and curing the barnacle bioadhesive interface remain unresolved. It is widely accepted that glands near the barnacle base plate periodically produce uncured cement that is distributed to the binding interface through a network of capillaries.8 Barnacle adhesive cures into a highly insoluble material, complicating biochemical analysis. Several cement proteins have been characterized and assigned roles in the cured adhesive. The multifunctional material that serves as adhesive is resistant to degradation, antibiotic, and selfhealing.16 The structure of the adhesive includes both intertwined fibers and unstructured domains.17−20 In studies of barnacle adhesive proteins, there is yet no evidence for the

ature employs a wide variety of adhesive strategies to enable temporary or permanent attachment to dry and wet surfaces. Geckos, spiders, and flies have evolved hierarchical nanostructures, scaling inversely with animal size, to facilitate locomotion and adhesion via van der Waals interactions,1 while gastropods have mucin gels with nonlinear rheological properties that modify film thickness and viscosity enabling snails to crawl up vertical surfaces.2 Means of permanent adhesion have evolved for a number of marine species. While mechanics may play a role, permanent underwater adhesion is frequently associated with proteinaceous composites where the proteins have significant fractions of post-translationally modified amino acid side chainsnotably modification of functional groups by hydroxylation or phosphorylation; both are found in mussel and sandcastle worm adhesive proteins.3−5 Hydroxylated tyrosine (3,4-dihydroxyphenyl-L-alanine, or Ldopa), in particular, has been implicated in the adhesion of mussels and many other organisms,3,6 but protein conformation and geometric factors may also strongly influence adhesion of L-dopa-containing biopolymers.7 © 2012 American Chemical Society

Received: April 25, 2012 Revised: June 18, 2012 Published: June 21, 2012 13364

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Image cone-beam reconstruction was performed using the Skyscan proprietary software. Ring artifact correction was applied, but only minimal smoothing and beam hardening corrections were applied. ImageJ (http://rsb.info.nih.gov/ij/) was used to stack and crop images, and OsiriX (www.osirix-viewer.com) was used to produce threedimensional volume renderings. Barnacle Reatttachment. After two to three additional weeks of growth, 4−7-mm-diameter barnacles were gently dislodged from the panels with cotton swabs, rinsed with distilled water, and placed onto clean 2 × 12.7-mm-diameter CaF2 windows and submerged in artificial seawater. For approximately three days (72 ± 8 h), the barnacles grew under the conditions outlined previously. This time period allowed for most barnacles to successfully adhere to the CaF2 substrate. Barnacles with cracks, chips, or other defects in the base plate were removed; the final analysis included 37 barnacles. Base plate growth was calculated by measuring base plate areas in optical micrographs taken immediately before and after the growth period.41 Optical Microscopy. Barnacle bases were monitored by bright field reflection and epifluorescence microscopy using a Nikon AZ100 microscope. Fluorescence microscopy utilized a green-fluorescent protein (GFP) filter cube (Nikon #96362) to distinguish autofluorescent regions of interest. This configuration was found to provide acceptable contrast, but may not have utilized the optimum wavelengths. Barnacles used for reattachment were imaged from below (e.g., their base plate) just prior to resettlement on CaF2 substrates and at the end of the ∼3 day growth period, after which barnacles had readhered. Time-lapse optical micrographs were also acquired of young adult barnacles grown from cyprids on glass microscope slides. For this, the slides were held at the tops of Petri dishes filled with artificial seawater containing Artemia. Slides were oriented so the barnacle bases were monitored through the air/slide interface, with the barnacle submerged in seawater. The light source was shuttered for fluorescence time-lapse imaging to minimize photobleaching. Confocal microscopy was performed using a Nikon Eclipse C1si system providing spectrally resolved fluorescence. Excitation was performed under 60× magnification with a 402 nm near-UV laser and emission detected between 420 and 800 nm with 10 nm spectral resolution. Shear Detachment Assay. The detachment assay was performed according to ASTM Standards D561842 with the following exception: Due to the very small forces required to detach the resettled barnacles (0.2−4 N), the suggested 4.5 N/s shear rate was not implemented. Each barnacle and CaF2 window were rinsed with distilled water, then secured vertically with an optical window mount. A digital force gauge (Shimpo FGV-1XY) attached to a hand crank was moved parallel to the CaF2 surface, making contact with the barnacle near the substrate/ adhesive interface. The peak force measured during detachment was measured by the force gauge. Because the mechanism is manually operated, care was taken to shear each barnacle from its substrate at the same rate, by moving the force gauge at a rate of approximately 1 mm/s. While this rate does not correspond to the 4.5 N/s outlined in ASTM D5618, effort was taken to ensure consistency between detachment events. To calculate critical detachment stress for each barnacle, we assumed the adhesion was localized to the change in area over 72 h, and divided the measured detachment force by the base plate area increase. Statistical Analysis. Both the BCS1 and the BCS(1 + 2) data sets were found to be normally distributed using the Anderson-Darling normality test,43 and an independent two-sample, two-tailed unequal variance t test showed that the average values are statistically different from each other (p < 0.001). Identical tests were performed for several other variables, including initial base plate area (p = 0.098), final base plate area (p = 0.085), change in base plate area (p = 0.58), growth period (p = 0.066), and amount of residual adhesive protein on the substrate after detachment (p < 0.1). None of the additional variables tested showed a significant difference between the two data sets, consistent with the inference that the difference in the critical shear stress was due to the presence of BCS2 in the adhesive layer. Characterization of the Residual Glue. Barnacles detached in the shear experiments described above left some adhesive on the CaF2

post-translationally modified amino acid L-3,4-dihydroxyphenylalanine (L-dopa) implicated in the adhesion of mussels and other marine organisms.21,22 Further, there is no consensus as to the specifics of the barnacle glue curing process, which has variously been linked to S−S bonding,22 quinone crosslinking,10,23 noncovalent bonding,24 and enzymatic clotting cascades;18 see Power et al.25 for a recent review. In some cases, cement proteins have been identified as surface or bulk functional,11,16,26,27 but otherwise, details of cement heterogeneity were beyond the scope of most studies. Most analyses of barnacle adhesive have not addressed variations in materials released to the interface. However, work reported by Saroyan et al.8 has made clear that barnacle adhesion is more complex than delivery of a single homogeneous secretion to the adhesive interface. Saroyan and co-workers examined the barnacle’s ability to repair a damaged adhesive interface and reattach to a new surface. They found that the concentric network of ducts secreted additional cement when the adhesive interface was compromised. This implied that the ducts were not clogged with cured cement. On the basis of optical microscopy observations, Saroyan et al. proposed that barnacles maintained unclogged ducts by a twostep process in which cement was first released from the ducts and then cleared out by a subsequent release of a flushing fluid. They also found that, when cement ducts were reused for reattachment or repair, the physical properties of the new adhesive often varied from that observed in original, undamaged interfaces. However, the histochemical and histological properties of the reattachment adhesive have been shown to be nearly identical.19,28 We note that the opaque, rubbery, and highly hydrated “gummy” cement29,30 found under some barnacles grown on silicones and in interstices under barnacles that have been slowly dislodged to leave a larger gap between base plate and the substratediffers substantially mechanically31 from well-cured adhesive and is beyond the scope of this investigation. To circumvent the analytical difficulties posed by the insolubility of barnacle cements and the challenge of observing processes occurring in a buried interface, we are employing in situ microscopies (e.g., optical, X-ray tomography,18,32 and vibrational spectroscopies33,34) to follow barnacle growth and development. In situ spectroscopies have also been employed by others to explore barnacle cyprid attachment35,36 and mussel adhesion.37,38 Here, we describe spatially and temporally resolved experiments that directly probe the barnacle bioadhesive interface as it develops, and demonstrate a twostep cementing process for barnacle adhesion.

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EXPERIMENTAL SECTION

Cyprid Settlement and Barnacle Maintenance. Barnacle cyprids (Balanus amphitrite) were settled onto glass microscope slides and 7.6 × 15.2 × 0.64 cm3 T2 silicone (Gelest) coated glass panels39 and reared as described by Holm et al.40 About 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 where they were submerged in artificial seawater (Instant Ocean dissolved in distilled water at 32 ppt, 25 °C, with ambient light). The seawater was changed three times a week immediately before each feeding. X-ray Tomography. The shell structure of a representative barnacle shell was imaged by X-ray microtomography using a Skyscan model 1172 tomography system with a 1.3 megapixel camera (Skyscan, Kontich, Belguim). The source X-ray voltage setting for scans was 80 kV with 0.5° rotation steps. Image voxel size was 9.9 μm. 13365

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structures associated with barnacle adhesive.8,45 Well-cured barnacle cement is typically semitransparent, and the interfacial and near-surface internal structures responsible for adhesion are visible in vivo when barnacles are grown on optically transparent substrates. Optical microscopy of the base plate is dominated by the calcified structures of the side plates and base plate, with faintly visible capillaries extending to the near surface region (Figure 2A). Compressing images acquired over

substrates. The residual glue was rinsed with distilled water, dried under N2 for approximately 24 h, and examined using transmission Fourier transform infrared (FTIR) spectroscopy with a Nicolet 750 spectrometer and a deuterated triglycine sulfate (DTGS) detector. Each sample was signal-averaged for 1024 scans at 4 cm−1 resolution with 2× zero filling and Happ-Genzel apodization. Reference spectra were acquired on CaF2 windows cleaned thoroughly with distilled water and Alkonox detergent. To better observe frequent differences among data sets, an absorption spectrum average was calculated from spectra normalized by the area under the amide I and II peaks. Atomic force microscopy (AFM) was performed in air using a Digital Instruments Nanoman operated in air. TESP cantilevers were used in intermittent contact mode to image the adhesive remaining after barnacle removal.

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RESULTS AND DISCUSSION Acorn barnacles like Balanus amphitrite attach permanently to intertidal surfaces by secreting adhesive that is protected by interlocked shell parietal (side) plates and base plate14,44 (Figure 1A). As the barnacle grows and enlarges its shell, it

Figure 2. (A) Bright field and (B) fluorescence micrographs of a portion of a barnacle basis edge, viewed from underneath the barnacle through an optically transparent substrate. In the fluorescence image, sections of concentric rings, contiguous with the capillary structures visible in both images, are observed due the presence of an autofluorescent material.

∼24 h into a time-lapse video (Supporting Information Video S1) of the base plate leading edge reveals that the capillaries appear at intervals a significant distance behind the growth front at the periphery. The same process observed with fluorescence microscopy reveals that the capillaries release an autofluorescent material to the interface, generating thin concentric bands (Figure 2B, Supporting Information Video S2). These observations suggest that development of the barnacle adhesive interface is accomplished by at least two distinct secretions: the first, Barnacle Cement Secretion 1 (BCS1), is released continually as the barnacle grows, and a second (BCS2) is released periodically (Figure 3A−C). The network of capillaries and ducts has historically been ascribed the role of primary adhesive delivery,8,16 but alternately could participate by repairing damaged adhesive by filling gaps beneath the barnacle, and as the source of adhesive for barnacle reattachment after displacement.8,46 While barnacle adhesive is known

Figure 1. (A) X-ray tomogram of an acorn barnacle, Balanus amphitrite, shell, cut away to reveal internal structures and (B) optical micrograph of adhesive remaining on a substrate after removal of barnacle soft tissue and shell.

lengthens its side plates and simultaneously expands to cover a larger area. Sequential growth involving molt cycles create a series of concentric rings or bands visible underneath the barnacle. The rings are formed by extension of the cuticular membrane44 followed by calcification; the adhesive is secreted and cured in place under the protection of the barnacle shell. The material remaining after removal of the barnacle’s soft tissue and calcareous structures (Figure 1B) exhibits both growth bands and remnants of the network of capillary 13366

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Figure 4. Diagonally split fluorescence micrographs of two barnacles, viewed from below through a transparent substrate, immediately after reattachment and three days later. Barnacles before (upper left portion) and after (lower right portion) the three day reattachment period exhibit lateral growth. Barnacle (B) secreted an autofluorescent material into the interface (white arrow) during the growth period, while barnacle (A) did not.

Figure 3. (A−C) Cutaway cartoon images of a barnacle, viewed from the side, depicting the order of events observed in the growth videos (Supporting Information). Between frames (A) and (B), the barnacle increases its diameter and secretes a proteinaceous material. Behind this growth front, a new capillary structure forms, and (C) secretes an autofluorescent material into the previously formed interface.

to be heterogeneous,16,21 the thin, buried interface of insoluble proteins has proved an obstacle to evaluating localized chemistries. In light of the time-lapse microscopy presented above, we hypothesized that both the initial secretion of BCS1 and the capillary duct secretion BCS2 play functional roles in adhesion. To test this hypothesis, shear adhesion test measurements were used to assess the contributions of the two components to adhesion. We reattached barnacles onto CaF2 optical substrates,39 and monitored their growth over a three-day period. During the growth period, each barnacle adhered to the substrate, but only a fraction secreted BCS2, as shown in fluorescent micrographs taken before and after the growth period (Figure 4A,B). We then detached the barnacles by applying a shear force parallel to the substrate/adhesive interface,42 recording the peak force during detachment. We calculated the critical shear stress for each detachment event and found that barnacles that secreted BCS2 adhered with approximately twice the strength, on average, of barnacles that did not (Figure 5), even after taking into account barnacle size, growth area, growth period, and several other variables (See Statistical Analysis Summary). The introduction of BCS2 into the interface is correlated with an increase in adhesion. Therefore, we find that barnacles adhere through a multistep process involving separate secretion of at least two optically distinct materials. The first observable secretion, BCS1, is an adhesive in its own right, while the autofluorescent BCS2 serves to enhance adhesion of the existing interface.

The shear detachment assay confirms that the initial secretion from the barnacle periphery, BCS1, acted as an adhesive. Detachment forces for cement containing BCS(1 + 2) agree reasonably well with previously reported barnacle adhesion strengths, which for smooth surfaces span from 80 kPa for Balanus crenatus on polymethylmethacrylate (PMMA) to 280 kPa for Balanus amphitrite on polyurethane;47 release experiments from reattached barnacles yielded similar values for rigid substrates and lower stresses for release from silicones.17,29,30,39,40,47,48 Here, the statistical hypothesis tests provide evidence that the autofluorescent BCS2 component in the barnacle adhesive is a major variable responsible for the difference in adhesive strengths. The sequential deposition and localization of barnacle adhesive secretions allows us to evaluate the structural and chemical nature of regions within the interface. Optical microscopy and atomic force microscopy (AFM) revealed the morphology of the residual adhesive is heterogeneous. Figure 6A shows a fluorescence micrograph of the residual adhesive from a reattached barnacle that reveals two very different BCS2 flow patterns. Near the region marked “1” is a partial remnant of a typical compact BCS2 band that connected into a continuous ring around the barnacle periphery. Discontinuities in the band are the result of fracture during detachment. In addition to this band, a different type of BCS2 spreading pattern is also observed from the next inner row of ducts. In this case, a series of spots are observed, each centered where a duct had been. The spot pattern is likely the result of a small 13367

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Figure 5. Box-whisker plot of stress (force per unit area, see methods section) to remove reattached barnacles from CaF2 substrates. Error bars denote the upper and lower quartiles of measurements, the box segments identify the middle two quartiles, and the sample mean is shown as a dot within the box. Barnacles secreting the autofluorescent material, BCS(1 + 2), to the interface showed significantly higher adhesion.

Figure 6. (A) Fluorescence micrograph of the adhesive remaining on the substrate from a barnacle that secreted fluorescent material from newly formed capillaries (arc), as well as previously formed capillaries (circular spots). Images (B−D) show AFM topography scans taken from corresponding regions (1−3) labeled in (A). The color gradient to the left of (C) corresponds to full scale height noted on each image. Panels (E,F) show optical micrograph (E) and AFM image (F) of adhesive remaining on substrate from a barnacle that secreted only BCS1. The inset images in (A) and (E) show the entire sample of residual cement (scale bar = 1 mm).

gap between the base plate and new substrate, disrupting the usual flow pattern. The continuous band from the newly formed region resembled the native barnacle interface. Adhesive regions that optically appear composed of mostly BCS2 (Figure 6A−D) as identified by fluorescence microscopy exhibited fibrillar structures, even in regions adjacent to brightly fluorescing areas. Adhesive containing solely BCS1 (Figure 6E,F) appeared nonfibrillar. Regions composed of a mixture of BCS(1 + 2) contained fibrils that intertwined, aggregating into increasingly larger diameter fibers (Figure 6B,D). Previous AFM studies report similar fibers on substrates after barnacle reattachment19 and on detached barnacle base plates.17,18,20 However, in these studies the fibrillar structures observed were not correlated to the secretory ducts distributed throughout the adhesive layer. The presence of BCS2 in the adhesive is correlated with increased fibril concentrations, and a mixture of both secretions resulted in a highly intertwined fibrillar network. The residual adhesive on the CaF2 windows was characterized using FTIR spectroscopy. Incorporating greater temporal control into the FTIR analysis revealed additional complexities not realized in previous bulk-scale FTIR analyses of barnacle cement.19,20,24,28,34,49 All IR spectra were consistent with proteinaceous secretions and no calcium carbonate was detected. On an individual basis, spectra of barnacle cement secretions from the same data sets often showed variations in biochemical structural components, complicating comparison of BCS1 and BCS(1 + 2). Therefore, our analysis focused on averaged data sets so that the most common differences between BCS1 and BCS(1 + 2) could be identified. We further

restricted analysis to the Amide I and II regions, which were the most readily interpretable for biochemical structure. Changes in the −CH, −NH, and −OH stretch and other regions were also observed, which will require further work to understand; the main features in the Amide regions are presented here. Using transmission FTIR spectroscopy, we find that the averaged, normalized amide IR absorption spectra of the two cements (Figure 7A) are similar but do show minor changes in line shapes, more easily distinguishable in the difference spectrum (Figure 7B). For barnacles that have secreted both fluids, IR absorptions at 1652, 1623, and 1514 cm−1 increase at the expense of the rest of the amide bands. The Amide I region reveals that the secondary structure after secretion of BCS2 is enriched in regions associated with random coil or α-helix (1652 cm−1) and cross-β motifs (1623 cm−1). The latter band is dominant and has previously been associated with amyloid50 and identified as a structural feature in deconvoluted spectra of barnacle adhesive.19,20,34 Amyloid is known as a high-strength biomaterial51−53 that has been found in other marine bioadhesives.54−56 In the Amide II region, the 1514 cm−1 peak is commonly associated with tyrosine side chains,57 although other phenolic compounds can also absorb in this region, suggesting that BCS2 contains a source of phenolic functional groups. This is further supported by histological studies which have demonstrated positive staining tests for phenolic compounds in barnacle cement.10,25,28,58,59 Other 13368

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Figure 8. Confocal optical micrograph (rendered from a series of vertical-stacked images, see Experimental Materials section) of a capillary orifice viewed from underside a live barnacle through a thin glass coverslip. The upper portion of the capillary (T structure) extends into the barnacle interior, while the orifice exits to the nearsurface region and fluorescence extends parallel to the periphery of the barnacle. The autofluorescent domains exhibit discontinuous brightness and submicrometer features. Color is rendered from peak emission intensity with 10 nm spectral binning.

adenine dinucleotide phosphate (NADPH), flavonoids, vitamins, and cross-linked tyrosine residues.65 The autofluorescent domains remain intact after the barnacle adhesive is treated by soaking in water, seawater, or ethylenediaminetetraacetic acid (EDTA) at pH 7. After exposure to 5% HCl, the blue autofluorescent regions in barnacle adhesive lose significant intensity; some intensity recovers upon exposure to a base (sodium bicarbonate). Conjugated and oxidized phenolic moieties are also known sources of autofluorescence.63,66−69 These include dityrosines found in elastic protein structures like resilin63,70 and from oxidative stress of proteins.69 Further, even in the absence of aromatic residues, intrinsic blue fluorescence in proteins and peptides can arise from secondary structures in amyloid-like fibrils.71,72 Such intrinsic fluorescence has been associated with water hydrogen-bonded to the protein backbone71 and delocalization of electrons in a cross-β structured protein backbone.72,73 We have observed components in barnacle adhesive consistent with both amyloid and phenolic groups; both can exhibit fluorescence that can be shifted or quenched by changes in pH.72,74 Our work demonstrates that the source of the blue autofluorescence is from within the barnacle capillary structures and penetrates through the interface to the surface. We note that autofluorescence has also been recently observed in sandcastle worm adhesive secretion organs and adhesive.68,75 Further studies are needed to identify the specific chemistry or structure(s) responsible for the autofluorescence of barnacle secretions, and their relation to adhesion strength and materials properties of the adhesive. The multicomponent interfaces created by barnacles form from two proteinaceous secretions that when combined nearly double the adhesion strength. The force per unit area needed to break an adhesive junction can be modeled using a fracture energy approach.76,77 The strength of the fractured joint is expected to follow the form given by

Figure 7. (A) Average normalized amide I and II bands of singlecomponent cement, BC1 (red trace), and two-component cement, BC(1 + 2) (blue trace). The absorption spectrum of the twocomponent adhesive has been shifted upward for clarity. (B) The difference spectrum of BC(1 + 2) − BC1 (1:1 scale). The difference spectrum was offset slightly as described in the text.

sessile marine organisms employ phenolic compounds, such as L-dopa in their adhesives. This modified tyrosine side chain is a key component implicated in cross-linking and adherence of marine mussel and sandcastle worm adhesives.3,4,60,61 While it has been reported that L-dopa is not a component in barnacle cement,21,22 other phenolic substituents could play similar roles and have been previously implicated in barnacle cement curing.10 Barnacle adhesive stains positive for peroxidase activity18 suggesting a biochemical pathway for phenolic cross-linking,28,62,63 although phenoloxidase inhibitors have not been demonstrated to prevent solidification of liquid cement.64 The FTIR and AFM results demonstrate differences in the morphology and biochemical structure of BCS1 and 2 when air-dried. Neither dry nor submerged environments perfectly represent the true physiological conditions of the adhesive interface under live barnacles. Further work is needed to determine how the morphology, biochemical structure, and other properties of BCS1 and 2 compare under different conditions including varying levels of hydration, salinity, and pH. Confocal optical microscopy of capillary orifices definitively reveals the capillaries as the source of autofluorescent material (Figure 8). Blue autofluorescent globular particles are released and accumulate into lateral bands extending circumferentially, and penetrating a micrometer or more through the prior interface formed during the secretion of BCS1, which exhibits a weak yellow, patchy autofluorescence. Blue autofluorescence in biological materials is common and is often associated with hydroxylated, phenolic compounds including reduced nicotinamide adenine dinucleotide (NADH) and nicotinamide

⎛ 2GW ⎞1/2 F ⎟ =⎜ A ⎝ t ⎠ 13369

([1])

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where F is the force to separate the surfaces, A is the contact area, G is the shear modulus of the adhesive, W the work of adhesion, and t the thickness of the adhesive layer. Similar fracture mechanics approaches are frequently applied to predict release and pull-off stresses for hard foulers like barnacles from compliant silicone coatings.77 In that case, the silicone coating is assumed to be the compliant component, as it is thick relative to the barnacle adhesive; we have also treated complexities of barnacle base plate compliance.32,78 For this exercise, we assume the substrate (CaF2) and barnacle shell are rigid relative to the proteinaceous adhesive. In our experiments, we observed a 2-fold increase in removal stress. For this magnitude of increase, the parameter GW/t would necessarily quadruple when BCS2 is secreted. Our AFM data do not indicate any change in thickness of the adhesive layer. Further, an evaluation of the FTIR data also supports the observation that there was no statistically significant difference in protein per unit area left on the surface after barnacle removal. Specifically, the relative density of protein in the residual adhesive was estimated by dividing the area under the amide bands by the corresponding base plate growth for each detachment event. The normalized absorbance for singlecomponent adhesive (BCS1 alone) and two-component adhesive BCS(1 + 2) are 5 ± 4 and 6 ± 6 arbitrary units/ mm2, respectively. Although the large variances for each set indicate a range of cohesive and adhesive failures within the data sets, the two measurements of residual protein density are not statistically different (p = 0.066). Therefore, we infer that either the interfacial material’s modulus or work of adhesion, or perhaps both, were substantially increased after BCS2 is secreted into the contact. Sun et al. have previously suggested a relationship between adhesive base plate modulus and release stress.31 Our results also demonstrate that the BCS2 secretion event occurs over a short time frame (hours) relative to the total time to develop one new “growth ring” (days). We also note that the stepwise increase in adhesion is correlated only with the secretion of BCS2, which as observed in epifluorescence micrographs and IR absorption spectra is in relatively low abundance in the cured adhesive interface. Combined, these observations suggest a disproportionate effect of BSC2 proteins and chemistries on adhesion relative to BCS1. Our experiments demonstrate that the increased adhesion results after the appearance of both amyloid-like fibrils and phenolic chemistries into, and through, a previously formed interfacial layer. The presence of functional amyloid can impart structural rigidity to the adhesive,51−53,79 while the phenolic residues may promote structural stability and adhesion through physical cross-linking within the protein network. The fibrils may also anchor the proteinaceous interface by conforming19 and coupling to the substrate, enhancing physical interactions with the surface. Both could explain the increase in adhesion observed through physical changes within the adhesive interface. Whether the secretion of BCS2 changes the structure of the interfacial proteins secreted earlier in the adhesion process is unknown. Side chain chemistry may also play a direct chemical role in adhesion. At least one recombinant protein identified in barnacle adhesive and characterized using sequencing and cloning may have calcite-binding affinity. However, it is present in small quantities in barnacle adhesive and has not been implicated in underwater attachment.80 It is possible that this protein has some affinity to adsorb to the CaF2 surface and plays a role in enhancing adhesion as well. Barnacles quite effectively adhere to a wide variety of surfaces,

and at least one other recombinant barnacle protein has been shown to have absorption affinity to surfaces with a wide range of characteristics, including positively and negatively charged, as well as hydrophobic surfaces.81 The phenolic components detected in BCS2 are definitely puzzling because their role in barnacle adhesion has been increasingly downplayed over the past 40 years. Scientists have long recognized the possibility for quinone or tanning chemistries in barnacle glue curing (see refs 10,23,28 and references therein), particularly because of their prevalence in structural membranes of arthropods and crustaceans. While oxidase chemistries necessary to accomplish these reactions have been implicated in barnacle adhesive curing,23 these ideas have been discounted due in large part to an inability of oxidase inhibitors in preventing solidification of barnacle secretions obtained from the capillary structures.64 Those experiments, however, did not address whether any other materials present in the interface were influenced by the inhibitors. Our experiments demonstrated proteinaceous adhesion before visible formation of capillary structures and their secretions. Our experiments point to a need to reexamine the role of phenolic components in barnacle interface secretions, particularly as they may contribute to chemical processes that stiffen the interface through intermolecular cross-linking. Notably, barnacles, more closely related on the evolutionary tree to arthropods (spiders and insects) than mollusks,82 have apparently not adapted costly posttranslational amino acid processing (such as conversion of tyrosine to L-dopa) in order to adhere to surfaces. Our experiments correlate the presence of proteins having β-sheet conformation and nanofibrillar structural motifs in an interface with improved adhesion. We also believe our results point to a need to revisit the role of phenolic components and enzymatic involvement in the curing process. How the barnacle creates a highly insoluble interface while maintaining the ability to reuse its network of ducts, perhaps through an additional “flushing” solution like that described by Saroyan,8 also remains an open question. The recognition of a multistep process for barnacle adhesion presents a paradigm shift that may open new directions to development of underwater adhesives,83 as well as instruct evaluation of the effectiveness of surface treatments targeting foulant adhesion through biochemistry.

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ASSOCIATED CONTENT

S Supporting Information *

Videos of time-lapse optical observations of barnacle interface development using optical microscopy and epifluorescence microscopy. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected], phone: (202) 767-5419. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We acknowledge support from the Office of Naval Research Coatings Program (Duke and NRL), and NRL Base 6.1 Funding. We enjoyed fruitful discussions with David Kidwell and John Russell. D.K.B. is a National Research Council Post Doctoral Associate. 13370

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