Bidirectional Surface Analysis of Monomolecular Membrane

Letter pubs.acs.org/NanoLett

Bidirectional Surface Analysis of Monomolecular Membrane Harboring Nanoscale Reversible Collapse Structures Amit K. Sachan and Hans-Joachim Galla* Institute of Biochemistry, Westfälische Wilhelms Universität, Wilhelm-Klemm-Strasse2, 48149 Münster, Germany S Supporting Information *

ABSTRACT: Determination of orientation of nanoscale collapse structures formed within a Langmuir film at the air−aqueous interface has not been possible by existing experimental techniques. This is however of special importance for pulmonary surfactant films, which form reversible surface-associated reservoirs (SARs) under dynamic lateral compression and expansion. The direction of these SARs with respect to the interface has hitherto remained uncertain. We designed a methodological approach to investigate the directionality of SARs formed in a functional analogue of the pulmonary surfactant lining, where we transferred the compressed film on a holey substrate and performed bidirectional surface imaging of the hole spanning monomolecular membrane harboring SARs. This unambiguously showed association of SARs with the membrane toward the air-side, in contrast to the up to now commonly accepted view of an orientation toward the aqueous phase. KEYWORDS: Atomic force microscopy, bidirectional surface imaging, collapse structure, pore-spanning monolayer, pulmonary surfactant, surface-associated reservoir

R

the directionality of the collapse formation and leaves an important question to assumption. Therefore, there is a need of either a technical advancement that can offer successful and reproducible studies of interfacial films at in situ with high lateral and vertical sensitivities or a methodological approach that can allow studying the individual collapsed structures associated with the interfacial membrane at ex situ in their in situ state. In this study, we have investigated the direction of the formation of reversible collapse structures in a functional analogue of pulmonary surfactant lining23,30,35 using a unique methodological approach. This method employs transfer of the compressed monomolecular film over a holey substrate (Figure 2a) with a condition that the substrate should allow performing bidirectional surface imaging of the membrane hovering over the holes to investigate the direction of the surface-associated collapse structures. Although interesting in concept, lipid monomolecular membrane spanning on the holes/pores and its stability has not been proven and documented so far. However, pore-spanning lipid bilayer studies have been reported,37,38 but none of those have employed the bidirectional surface imaging of the membrane. For bidirectional imaging, a technical difficulty lies in the thickness of the chosen porous substrate so far. Any holey/porous substrate of less than 100 nm thickness, depending on the hole size, can provide an alternative for the investigation but it may not be stable by its own, therefore, supporting such holey substrate on another mesh-like solid substrate can bring forth a requisite support for the bidirectional

ecently, tremendous efforts have been made in understanding the process of formation of three-dimensional collapse structures during the compression of Langmuir films, especially of biological importance. 1−8 The pulmonary surfactant (natural, modified-natural or synthetic) films, containing surfactant specific proteins B (SP-B) and/or surfactant specific proteins C (SP-C), exhibit a novel reversible mechanism of collapse at the air−aqueous interface on the way to neutralize the surface tension of the aqueous subphase and form surfaceassociated reservoirs (SARs) or collapse structures, which quickly respread in the interfacial layer at the increase of surface tension.9−12 The existence of the same surfactant efficient mechanism has been proposed in the native pulmonary surfactant lining present at the air−alveolar interface for a mechanically efficient continuous breathing process in the lungs,13−17 which observe continuous surface tension variations between 30 and 0 mN/m at the air−alveolar interface.18,19 However, in this mechanism the direction of the formation of these reservoir structures with respect to the interface has remained uncertain even after decades of intense research.20−32 A number of in situ techniques28,29,33,34 have been employed for the purpose, but thus far none of those are able to provide highresolution states of individual nanoscale collapses. For highresolution investigations, compressed films have been primarily deposited on the solid supports and studied via electron microscopy35,36 and atomic force microscopy8−10,19,23,35 at ex situ (Figure 1a−d). As the deposition process on the solid support can lead to similar perceptible topography of the collapse structures formed in either direction (i.e., air-side or aqueous-side) of the interface due to the possible surface layer rearrangements within the collapsed structures (Figure 1e), this approach of high-resolution investigation also denies revealing © 2013 American Chemical Society

Received: October 25, 2012 Revised: January 30, 2013 Published: February 7, 2013 961

dx.doi.org/10.1021/nl303928m | Nano Lett. 2013, 13, 961−966

Nano Letters

Letter

Figure 1. Typical AFM topography images of a functional analogue (1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC):1,2-dipalmitoyl-sn-glycero-3phosphoglycerol (DPPG):SP-C mixture) of pulmonary surfactant lining, compressed to 65 mN/m surface pressure and transferred on mica via horizontal hydrophilic transfer technique. Images a and b represent the pattern of multilayered surface-associated reservoirs (SARs) or collapsed structures (bright network) associated with the background monomolecular membrane, (z-range 0−90 nm). Images c and d emphasize the geometry of individual collapsed structures at 65 mN/m, (z-range 0−70 nm). Bright colors within each AFM topography image represent higher values of the signal. Image e demonstrates possible lipid layer rearrangements within collapsed structures during deposition on a solid substrate and formation of undifferentiable topography under two different possible directions of formation of collapse with respect to the interface.

Figure 2. An alternative solid support and transfer of the compressed monomolecular membrane. A proposed holey/porous substrate (a) for the transfer of the compressed membrane to investigate the orientation of the collapsed structures through bidirectional surface analysis of the monomolecular membrane hovering over the holes/pores. (b and c) Scanning electron microscopy (SEM) images of Quantifoil holey carbon film spanned over an EM grid, an alternative for the proposed substrate. Transfer of compressed monomolecular membrane on the holey carbon substrate, followed by coating (10 nm amorphous carbon) from topside, reveals covering of some of the holes in the SEM image (d). SEM imaging at an angle (e and f) shows steps over the covered holes. The white box in f focuses the area of steps of collapsed structures. AFM topography (g, z-range 0−850 nm) also shows covering of some of the holes by the transferred membrane, and the differences in penetration of the cantilever tip are shown in cross section profiles of uncovered and covered holes (h and i), respectively. 962


Nano Letters

Letter

Figure 3. Envisioned observable topography during bidirectional surface study via AFM of monomolecular membrane hovering over the holes and harboring collapsed structures in either of two possible directions, that is, toward air-side (a) or toward water-side (b). (c) An illustration of the effect of the curvature of area around an object in the measured topography of the object by cantilever tip in AFM. h, h′, and h″ depict three different observable heights of the same height object positioned at different curvature areas.

SEM imaging was performed at an angle that led us to observe a few structural steps on the membrane transferred over the holes (Figure 2e and f), which hints the presence of collapsed structures over the holes. Atomic force microscopy (AFM) topography imaging at the corresponding places in the mesh also reproduced these results. Both the covered holes and the uncovered ones can be readily identified based on cantilever tip penetration (Figure 2g). The tip shows penetration into the uncovered holes by ∼700−800 nm (Figure 2h), while the covered holes do not allow such penetration.40 The cross section analysis across the covered holes presents curvature in the membrane transferred over the holes, and the depth of this curvature lies between 30 and 70 nm at the center. Besides, the cross section profile also reveals structural steps over the covered holes (Figure 2i), which suggests the presence of collapsed structures over the monomolecular membrane toward the air-side and requires further in detail examination. Bidirectional Correlative SEM-AFM Surface Imaging of Monomolecular Membrane. The finding of collapsed structures associated with the membrane toward the air phase was further examined by the bidirectional correlative SEMAFM surface study of the individual covered holes. For this study, at each set of experiments at least two holey Quantifoil grids were used at a time during the transfer of the compressed film. Around 30 min after transfer, one grid was placed upsidedown to preserve the collapsed structures from any structural rearrangements due to the gravity, if collapse structures form toward the air phase. Thereafter, these grids with “upside” and

surface accessibility of the samples lying over the holes. To test the proposal, we chose Quantifoil holey carbon films with thickness of ∼20 nm, regular holes of 1 μm size and hole spacing of 0.6 μm, spanned on electron microscopy (EM) grids, as an alternative substrate for our study (Figure 2b and c). The size of the holes was chosen as per the observed width of the collapsed structures, to accommodate the collapsed structures over the individual holes (Figure 1c and d). Transfer of the Compressed Membrane on a Holey Support. During transfer of the compressed film on a substrate by the Langmuir−Blodgett technique, a lateral structural shift of the collapsed structures can occur due to the high lateral stress associated during the vertical deposition of the collapse structures. Therefore, we performed the horizontal hydrophilic transfer of the compressed film (surface pressure 65 mN/m) on the holey substrate using house-made transfer module.39 After the transfer, without any verification the grids were coated on the side of film transfer by either 10 nm amorphous carbon (C) or 2 nm C + 1.5 nm Pt/C. The aim of the coating was to provide mechanical stability and longevity to the monomolecular membrane, if spanning on the holes, and to avoid membrane damage during subsequent imaging of the respective areas. At first, the successful transfer of the compressed membrane on the grids was verified by scanning electron microscopy (SEM) at the side of membrane transfer. We observed covering of some of the holes of the grids partially and/or completely by the membrane transfer (Figure 2d). The fraction of the holes covered within a mesh of a grid varied from 0 to 10%. Further, 963


Nano Letters

Letter

Figure 4. Bidirectional correlative SEM-AFM surface imaging of coated (2 nm C + 1.5 nm Pt/C) monomolecular membrane hovering over the holes. Images a−f belong to the topside-coated sample and images g−l to the backside-coated sample. The coated side of the membrane over the holes in each sample does not produce contrast in SEM images (a and h); however, the uncoated side provides contrast (b and g) and differentiates areas of collapsed regions (dark areas) from the background monomolecular membrane over the hole. Darker areas over the holes in these SEM images (b and g) represent indirectly the higher height of the collapsed structures. In topside-coated membrane, the topside topology result by AFM shows a curved collapsed structure over the hole (c, z-range 0−120 nm), while backside topology (d, z-range 0−55 nm) provides a curved membrane with a higher curvature place, which overlaps well with the place of collapsed structure observed at the other side of the membrane. In the backside-coated membrane, the topside topology by AFM shows clear structures of collapse (i, z-range 0−115 nm), while the backside yields a curved membrane with no collapsed structure (j, z-range 0−55 nm). The areas of collapsed structures observed in AFM images correlate absolutely with the dark areas observed in SEM images. Bright colors within the AFM height images represent the higher values of the signal. Amplitude signal images (e and f, k and l) provide better distinction of structures over the hole. Scale bars, 500 nm.

“upside-down” conditions were coated by 2 nm C + 1.5 nm Pt/C and termed as topside-coated and backside-coated, respectively. For the bidirectional AFM topography imaging, a specialized house-made grid-supporting holder was designed to avoid the sample damage by surface contact of the other side of the grid with the support while imaging at one side. Under two possible directions of formation of SARs and with added effects of two different directions of coating, we need to envision following topography scenarios for the bidirectional surface imaging of the membrane lying over the holes. Condition I is if collapse structures form toward the air phase (Figure 3a). Topside topology may not reveal clear steps of collapsed structures over the holes if coating is performed on the topside, as coating material can push the structures down on the curved monomolecular membrane, which will lead to topology artifacts due to the curvature effect during AFM investigation (Figure 3c), and the backside topography should show a curved membrane with higher curvature at the places of collapsed structures lying on the other side, that is, topside. However if coating is performed on the backside, topside topology should yield clear structural steps of collapse as coating will relax the curvature of the hanging membrane, and the backside imaging should observe an almost flat membrane. Condition II is if collapse structures form toward the aqueous

phase (Figure 3b). Topside surface imaging should show a curved membrane without any structure over the hole, and imaging at backside should yield clear structural steps, when coating is performed at the topside. However, if coating is performed at the backside, topside surface imaging should again show a curved membrane without any structural step while backside imaging should observe structures with affected step heights. Other than this, if collapse structures form toward both sides of the interface, mixed topography results are expected. Figure 4 presents the bidirectional correlative SEM-AFM surface imaging results of the compressed monomolecular membrane spanning over the holes. SEM imaging on the coating side of each grid shows almost no structural information about the collapsed structures over the holes (Figure 4a and h), whereas the other side of the coating side yields contrast over the holes, where the dark regions over the hole represent the areas of more than a monomolecular lipid membrane and demarcate the area of collapsed structures from the background monomolecular membrane (Figure 4b and g). Correlative AFM bidirectional surface imaging results support this (Figure 4c−f and i−l). We observe curved collapsed structures with minor visible steps from topside and a curved membrane with a higher curvature place during backside imaging, in the topside-coated samples. The higher curvature place in the backside topography 964


Nano Letters

Letter

structural steps in the multilayered collapsed structure associated on the topside of the monomolecular membrane hovering over the hole. In the light of this finding, we undoubtedly prove the formation of the reversible surface-associated structures in a compressed synthetic pulmonary surfactant membrane in the airphase and rule out the conventional belief of formation of such structures toward the aqueous-phase. Moreover, the layered structure of the collapses over the holes appears to represent close to the in situ state of these structures. The next question arises hereafter about how these layered collapse structures form in air. Therefore, we propose a mechanism of formation of these layered structures at in situ (Figure 6). When a

correlates well with the place of collapse structure observed in the topside topography (Figure 4c and d, and Supporting Information, Figure S1). However, in the backside-coated samples we observe clear structural steps from topside over the holes and almost flat surface in backside surface imaging (Figure 4i and j, and Supporting Information, Figure S2). These results match finely with the predictions for condition I with coating effects and confirm the formation of collapsed structures in the air-phase. Furthermore, to avoid any unnoticed artifacts due to the coating, we attempted to image these collapsed structures in their native state on the hanging monomolecular membrane without any coating. We understand that the presence of the collapsed structure on the hole-spanning monomolecular membrane may provide vulnerability as well as stability to the hanging membrane depending on its position with respect to the center of the hole, amount of the collapsed material or weight, and its continuity across the hole. Therefore, maximum care was taken to provide shock-free handling to the freshly transferred grids. First, we performed SEM imaging of the grid at low magnification and at low current to observe covered holes as well as to avoid maximum possible electron beam damage to the uncoated and unsupported monomolecular membrane (Figure 5a). Later on, we performed a correlative AFM

Figure 6. Schematic of the proposed mechanism of formation of collapsed structures with bilayer stacks toward air-side. Twodimensional cross section view (not up to the scale).

monomolecular film at the interface (Figure 6, step 1) is compressed beyond its equilibrium surface pressure, nanoscale nucleating inverse bilayer structures form and protrude toward the air-side from the favorable phase (such as fluid phase) of the interfacial film (Figure 6, step 2). These protrusions grow to certain heights under lateral compression (Figure 6, step 3) and subsequently either fall freely at their places forming the multilayered structures with the heightened collapse structure or bend and fall on the nearby protrusions leading to the formation of multilayered structures (Figure 6, step 4; Figure 1c and d). Preassuming the collapse formation toward the airphase, a similar mechanism has been proposed previously in other Langmuir films.36,41,42 Our approach clears an important question related to the formation of surface-associated reservoirs during the compression of the pulmonary surfactants. Having a clear idea about the directionality of the collapse structures will absolutely pave way to establish a clear mechanism and further in the understanding of the roles of surfactant-specific proteins in the formation of these reversible collapse structures. In general, our approach provides a better understanding of industrially and biologically

Figure 5. Topside investigation of uncoated unsupported monomolecular membrane hovering over the holes. SEM image (a) shows partially and completely covered holes by the transferred membrane. The blue box in the SEM image indicates the correlative area of AFM topography (b, z-range 0−850 nm). AFM topography images over the individual covered holes clearly show presence of collapsed structures toward air-side (c and d, z-range 0−115 nm). Cross section profile (e) across the topography image (d) depicts multilayer nature of collapsed structure.

surface study of the hole-spanning force-sensitive monomolecular membrane, which again revealed the collapse structures in the topside topography (Figure 5b−d). The cross section profile (Figure 5e) across the topography image (Figure 5d) shows clear 965


Nano Letters

Letter

relevant Langmuir films’ three-dimensional structures. Moreover, our unique methodology of bidirectional surface analysis offers immense potential in the area of membrane biophysics and material science.

■

(21) Schürch, S.; Bachofen, H.; Possmayer, F. In Complexity in Structure and Function of the Lung; Hlastala, M. P., Robertson, H. T., Eds.; Marcel Dekker: New York, 1998; pp 35− 74. (22) Schürch, S.; Green, F. H. Y.; Bachofen, H. Biochim. Biophys. Acta 1998, 1408, 180−202. (23) Amrein, M.; Nahmen, A. V.; Sieber, M. Eur. Biophys. J. 1997, 26, 349−357. (24) Knebel, D.; Sieber, M.; Reichelt, R.; Galla, H. J.; Amrein, M. Biophys. J. 2002, 82, 474−480. (25) Baoukina, S.; Tieleman, D. P. Biophys. J. 2011, 100, 1678−1687. (26) Casals, C.; Cañadas, O. Biochim. Biophys. Acta 2012, 1818, 2550−2562. (27) Zuo, Y. Y.; Tadayyon, S. M.; Keating, E.; Zhao, L.; Veldhuizen, R. A. W.; Peterson, N. O.; Amrein, M.; Possmayer, F. Biophys. J. 2008, 95, 2779−2791. (28) Schief, W. R.; Antia, M.; Discher, B. M.; Hall, S. B.; Vogel, V. Biophys. J. 2003, 84, 3792−3806. (29) Takamoto, D. Y.; Lipp, M. M.; Nahmen, A. V.; Lee, K. Y. C.; Waring, A. J.; Zasadzinski, J. A. Biophys. J. 2001, 81, 153−169. (30) Krol, S.; Janshoff, A.; Ross, M.; Galla, H. J. Phys. Chem. Chem. Phys. 2000, 2, 4586−4593. (31) Wang, L.; Cai, P.; Galla, H. J.; He, H.; Flach, C. R.; Mendelsohn, R. Eur. Biophys. J. 2005, 34, 243−254. (32) Baumgart, F.; Ospina, O. L.; Mingarro, I.; Crespo, I. R.; PérezGil, J. Biophys. J. 2010, 99, 3234−3243. (33) Lipp, M. M.; Lee, K. Y. C.; Zasadzinski, J. A.; Waring, A. J. Science 1996, 273, 1196−1199. (34) Follows, D.; Tiberg, F.; Thomas, R. K.; Larsson, M. Biochim. Biophys. Acta 2007, 1768, 228−235. (35) Sachan, A. K.; Harishchandra, R. K.; Bantz, C.; Maskos, M.; Reichelt, R.; Galla, H. J. ACS Nano 2012, 6, 1677−1687. (36) Ries, H. E. Nature 1979, 281, 287−289. (37) Mey, I.; Stephan, M.; Schmitt, E. K.; Müller, M. M.; Amar, M. B.; Steinem, C.; Janshoff, A. J. Am. Chem. Soc 2009, 131, 7031−7039. (38) Steltenkamp, S.; Müller, M. M.; Deserno, M.; Hennesthal, C.; Steinem, C.; Janshoff, A. Biophys. J. 2006, 91, 217−226. (39) Dutta, A.; Kundu, S.; Sanyal, M. K.; Daillant, J.; Luzet, D.; Blot, C.; Struth, B. Phys. Rev. E 2005, 71 (041604), 1−7. (40) It is important to point out that the tip shape and geometry and the applied imaging force will determine the maximum penetration depth achieved by the cantilever tip into a hole of particular size. (41) Birdi, K. S.; Vu, D. T. Langmuir 1994, 10, 623−625. (42) Chen, X.; Wiehle, S.; Weygand, M.; Brezesinski, G.; Klenz, U.; Galla, H. J.; Fuchs, H.; Haufe, G.; Chi, L. J. Phys. Chem. B 2005, 109, 19866−19875.

ASSOCIATED CONTENT

S Supporting Information *

Experimental details and Figure S1 and Figure S2. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*Tel.: +49 251 8333200. Fax: +49 251 8333206. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We would like to thank Institute of Medical Physics and Biophysics (IMPB), Münster for financial support as well as availing the instruments to A. K. Sachan. We thank U. Keller and H. Nüsse, IMPB for technical assistance, and S. Tacke, IMPB for helpful discussions. A.K.S. conceived the complete idea, designed and performed all sets of experiments, and completed the analysis. H.J.G. critically assessed the results. H.J.G. and A.K.S. wrote the manuscript.

■

REFERENCES

(1) Lee, K. Y. C. Annu. Rev. Phys. Chem. 2008, 59, 771−791. (2) Baoukina, S.; Monticelli, L.; Risselada, H. J.; Marrink, S. J.; Tieleman, D. P. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 10803−10808. (3) Lorenz, C. D.; Travesset, A. Langmuir 2006, 22, 10016−10024. (4) Gopal, A.; Lee, K. Y. C. J. Phys. Chem. B 2001, 105, 10348− 10354. (5) Ybert, C.; Lu, W.; Möller, G.; Knobler, C. M. J. Phys. Chem. B 2002, 106, 2004−2008. (6) Baoukina, S.; Monticelli, L.; Amrein, M.; Tieleman, D. P. Biophys. J. 2007, 93, 3775−3782. (7) Bron, A. J.; Tiffany, J. M.; Gouveia, S. M.; Yokoi, N.; Voon, L. W. Exp. Eye Res. 2004, 78, 347−360. (8) Rantamäki, A. H.; Telenius, J.; Koivuniemi, A.; Vattulainen, I.; Holopainen, J. M. Prog. Retinal Eye Res. 2011, 30, 204−215. (9) Alonso, C.; Alig, T.; Yoon, J.; Bringezu, F.; Warriner, H.; Zasadzinski, J. A. Biophys. J. 2004, 87, 4188−4202. (10) Zhang, H.; Fan, Q.; Wang, Y. E.; Neal, C. R.; Zuo, Y. Y. Biochim. Biophys. Acta 2011, 1808, 1832−1842. (11) Jiao, X.; Keating, E.; Tadayyon, S.; Possmayer, F.; Zuo, Y. Y.; Veldhuizen, R. A. W. Biophys. Chem. 2011, 158, 119−125. (12) Zhang, H.; Wang, Y. E.; Fan, Q.; Zuo, Y. Y. Langmuir 2011, 27, 8351−8358. (13) Rugonyi, S.; Biswas, S. C.; Hall, S. B. Respir. Physiol. Neurobiol. 2008, 163, 244−255. (14) Possmayer, F.; Hall, S. B.; Haller, T.; Peterson, N. O.; Zuo, Y. Y.; Serna, J. B.; Postle, A. D.; Veldhuizen, R. A. W.; Orgeig, S. Respir. Physiol. Neurobiol. 2010, 173S, S55−S64. (15) Pérez-Gil, J. Biochim. Biophys. Acta 2008, 1778, 1676−1695. (16) Zuo, Y. Y.; Veldhuizen, R. A. W.; Neumann, A. W.; Peterson, N. O.; Possmayer, F. Biochim. Biophys. Acta 2008, 1778, 1947−1977. (17) Wüstneck, R.; Pérez-Gil, J.; Wüstneck, N.; Cruz, A.; Fainerman, V. B.; Pison, U. Adv. Colloid Interface Sci. 2005, 117, 33−58. (18) Schürch, S.; Goerke, J.; Clements, J. A. Proc. Natl. Acad. Sci. U.S.A. 1978, 75, 3417−3421. (19) Schürch, S.; Bachofen, H.; Weibel, E. R. Respir. Physiol. 1985, 62, 31−45. (20) Bachofen, H.; Gerber, U.; Gehr, P.; Amrein, M.; Schürch, S. Biochim. Biophys. Acta 2005, 1720, 59−72. 966


Bidirectional Surface Analysis of Monomolecular Membrane

Recommend Documents