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Preliminary analysis of polyhydroxyalkanoate inclusions using atomic force microscopy
FEMS Microbiology Letters 226 (2003) 113^119
www.fems-microbiology.org
Preliminary analysis of polyhydroxyalkanoate inclusions using atomic force microscopy Douglas Dennis a; , Caroline Liebig b , Tara Holley b , Kara S. Thomas b , Amit Khosla b , Douglas Wilson b , Brian Augustine c a
Department of Life Sciences, Arizona State University West, P.O. Box 37100, Phoenix, AZ 85069-7100, USA b Department of Biology, James Madison University, Harrisonburg, VA, USA c Department of Chemistry, James Madison University, Harrisonburg, VA, USA Received 5 May 2003 ; received in revised form 8 July 2003 ; accepted 15 July 2003 First published online 26 August 2003
Abstract Atomic force microscopy analysis of polyhydroxyalkanoate (PHA) inclusions isolated from sonicated Ralstonia eutropha cells revealed that they exhibit two types of surface structure and shape; rough and ovoid, or smooth and spherical. Smooth inclusions possessed linear surface structures that were in parallel arrays with 7-nm spacing. Occasionally, cracks or fissures could be seen on the surface of the rough inclusions, which allowed a measurement of approximately 4 nm for the thickness of the boundary layer. When the rough inclusions were imaged at higher resolution, globular structures, 35 nm in diameter, having a central pore could be seen. These globular structures were connected by a network of 4-nm-wide linear structures. When the inclusions were treated with sodium lauryl sulfate, the boundary layer of the inclusion deteriorated in a manner that would be consistent with a lipid envelope. When the boundary layer was largely gone, 35nm globular disks could be imaged laying on the surface of the filter beside the inclusions. These data have facilitated the development of a preliminary model for PHA inclusion structure that is more advanced than previous models. 7 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords : Atomic force microscopy; Poly-3-hydroxyalkanoate; Inclusion
1. Introduction Bacterial polyhydroxyalkanoates (PHAs) are biodegradable polyesters that are found as intracellular inclusions that are generally between 200 and 500 nm in diameter [1,2]. At present, very little is known about their structure and biogenesis. In Ralstonia eutropha, three proteins have been shown to exist at their surface, PhaC, PhaP and PhaR. PhaC is the PHA synthase and is responsible for the formation of the polyester from 3-hydroxy fatty acylCoA molecules [2,3]. PhaP is thought to be a structural protein because it is found in very large quantities in PHA-accumulating cells and has profound e¡ects on the shape and number of PHA inclusions when it is deleted or
* Corresponding author. Tel. : +1 (602) 543 6934; Fax : +1 (602) 543 6073. E-mail address : [email protected] (D. Dennis).
overexpressed [4]. PhaR regulates the expression of PhaP [5,6]. Early electron microscopic studies (carbon replica) on inclusions from Bacillus megaterium supported the existence of a membrane layer at the surface of the inclusion [7], but later studies on inclusions from Pseudomonad species imaged a regular lattice-like surface architecture that is reminiscent of a bacterial S-layer [8^12]. Because of this, some researchers suggested that a protein lattice covers the inclusions, most logically comprised of PhaP. The possibility of whether the boundary layer is either membrane or protein has been somewhat clari¢ed in that researchers have determined that boundary layer thickness is 4 nm [13,14], a fact that would seem to exclude the possibility of a lipid bilayer surrounding the inclusion because lipid bilayers are approximately 8 nm in thickness. Mayer and Hoppert have o¡ered strong theoretical arguments based on hydrophobic interactions of membranes as to why this boundary layer is most probably a lipid monolayer. Nonetheless, de¢nitive evidence on the composition of the boundary layer and the arrangement of proteins at its sur-
0378-1097 / 03 / $22.00 7 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/S0378-1097(03)00610-4
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face is scarce, and several models of inclusion structure continue to exist [2,8,9,11,12]. Recently, atomic force microscopy (AFM) studies on inclusions from Comomonas acidovorans have given support to the existence of an envelope around the inclusion [15]. The possible existence of a lipid boundary layer is quite interesting because a corollary implication is that there would have to be structures that would allow for the ingress of substrates for polymerization, as well as egress of depolymerized monomers to be released into the cytoplasm. The recent AFM work also noted the presence of uncharacterized 30-nm ‘globular’ structures on the surface of the inclusions. The use of AFM to analyze PHA inclusions surfaces is particularly appropriate because of the minimal amount of preparation necessary to image the inclusions. Past inclusion models su¡er from the fact that they were developed from electron micrographs of PHA inclusions that required signi¢cant sample preparation and manipulation prior to microscopy, possibly disrupting the integrity of the inclusion surface. In this study, we present AFM images that con¢rm the existence of the boundary layer and which reveal a network of organized structures on the surface of the inclusion that may function in structure and metabolism, as well as facilitating movement of molecules through the boundary layer.
2. Materials and methods 2.1. Bacterial strains and culture conditions R. eutropha was grown in Luria broth containing no NaCl (LBN) for routine growth and was maintained on LBN agar. For induction of PHA accumulation the bacteria was grown in Schlegel’s nitrogen-limited minimal media containing 0.1% (w/v) ammonium sulfate and 0.5% fructose [16]. 2.2. Inclusion preparation R. eutropha cultures were harvested when inclusions were clearly visible intracellularly (light microscopy), which was usually about 24 h post-inoculation when the Schlegel’s nitrogen-limited media was used. Five microliters of culture was added to 5 ml of 50 mM Tris (pH 8.0)^ 1 mM EDTA (or 50 mM Tris (pH 8.0)^0.15 M NaCl) and the suspension was added to a Falcon 2059 polypropylene tube and sonicated using an Tekmar 400 watt sonicator with a mid-sized probe. Sonication was done in an ice bath for 5 min using 5-s-on and 7-s-o¡ cycles. To rid the sonicated suspension of intact cells, the tube was centrifuged at 1000Ug for 10 min at 4‡C. The top 4 ml of the supernate was removed to a new tube and placed on ice. For AFM studies, varying amounts of this suspension were added to 10 ml of 50 mM Tris (pH 8.0)^1 mM EDTA and this was
¢ltered through a 250-nm pore size polycarbonate ¢lter. After the initial 10 ml, the ¢lter was washed with 2U5 ml of TE, followed by 5 ml of water. The ¢lter was immediately removed, adhered to a metal puck (12 mm; Ted Pella, Inc) and was directly imaged by AFM. 2.3. Sodium dodecyl sulfate (SDS)-mediated disintegration of boundary layer This procedure was the same as above, except that the 4 ml of supernate removed after the 1000Ug centrifugation was now added to 36 ml of a prewarmed (45‡C) SDS solution such that the ¢nal SDS concentration was 0.5% (w/v). The suspension was gently mixed and incubated at 45‡C for 5 min. Varying amounts of the suspension were removed and ¢ltered through 250-nm pore size polycarbonate ¢lters, followed by two 5-ml washes with deionized water. Filters were processed as above. 2.4. AFM AFM was performed on a Digital Instruments Multimode instrument operating in tapping mode to acquire both height and phase images simultaneously. All imaging was performed in air at room temperature using silicon tapping mode probes manufactured by Olympus and purchased from Digital Instruments. Typical resonance frequencies for the probes were V250^300 kHz. Phase imaging was obtained by applying a relatively large tapping force (low setpoint) to the surface of the inclusion bodies. Typical scan speeds were 0.3^0.7 Hz.
3. Results In general, two types of inclusions (Fig. 1A,B.) were observed by AFM. The majority of inclusions tended to be ovoid in appearance and have a ‘rough’ or ‘bumpy’ surface. Occasionally, another type of inclusion that is rounder and had a smoother surface was seen. The ‘smooth’ inclusions had a di¡erent pro¢le in that they are more ‘mushroom-like’ and did not extend as high above the surface of the ¢lter (cross-section data not shown). Generally, their height pro¢le was about half of the pro¢le of rough inclusions. The proportion of smooth inclusions in the preparation increased if, 1) bacterial cells stored at 4‡C for several days were used, 2) the cells were subjected to longer periods of sonication, or 3) higher levels of EDTA were used (5 or 10 mM). When the ¢lters containing inclusions were stored at room temperature for a week and re-imaged, wide ¢ssures or cracks on the surface of the inclusion could be seen, presumably due to drying. The surface of the sublayer at the bottom of these ¢ssures is consistently 4 nm below the surface of the ‘rough’ inclusion (cross-section analysis, data now shown).
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Fig. 1. AFM images of PHA inclusions. AFM height (A) and phase (B) images of two types of PHA inclusions isolated from sonicated cells. In the height image the brightness of the image is directly related to the height of the inclusion. Terracing at the edge of the inclusion in the phase image is an artifact and should be disregarded. C: AFM high resolution phase image of the surface of a smooth inclusion showing 7-nm parallel arrays. D : AFM phase image of rough inclusion showing 35-nm globular structures with central pore.
3.1. High resolution studies When the ‘rough’ and ‘smooth’ inclusions were imaged using AFM at resolutions near the limit of the instrument, a di¡erent type of structure could be identi¢ed on the surface of the inclusion. The smooth inclusions had linear strands in parallel arrays traversing the surface of the inclusions (Fig. 1C). These parallel arrays did not extend uninterrupted across the surface of the inclusions, but could be seen traversing the surface of the inclusion in regions that were then interrupted by another set of linear arrays traversing a di¡erent direction. It would be analogous to the strokes of a paintbrush across the surface of the inclusion. The distance between the lines of the parallel
array was quite consistent and was measured by cross-sectional analysis at 7 nm (data not shown). The ‘rough’ inclusions exhibit quite a di¡erent surface morphology. The surfaces of these inclusions were populated with globular structures that were roughly spherical and which had a central pore (Fig. 1D). The globular structures were consistent in size, approximately 35 nm in diameter. Cross-sectional analyses of these structures revealed that the average pore diameter of the structure was about 15 nm (at the surface). The collar of the structure was about 12^15 nm in diameter and extended 4^12 nm above the surface of the inclusion (data not shown). Generally, many pores can be seen and they appear to be connected by linear structures that traverse the surface of the inclusion,
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Fig. 2. High resolution AFM phase image of PHA inclusion. Speci¢c structures mentioned in text: 1: Parallel arrays with 7-nm spacing; 2: linear network-like structures ; 3: porin-like structures with central pore. Network structures appear to be more prominent at the edge of scan because of tip convolution.
making it appear that there is a cytoskeletal structure encompassing the inclusion (Fig. 2). The average width of these structures is about 4 nm, but they can be wider, particularly at the points that they connect to the ‘porin-
like’ structures. In the highest resolution images, the same parallel linear arrays (7 nm spacing) seen on smooth inclusions could be imaged beneath the cytoskeletal-like network (Fig. 2).
Fig. 3. AFM phase images of inclusions treated with warmed 0.5% SDS. The left image shows the boundary layer in earlier stages of deterioration, where the right image shows an inclusion in the later stage of deterioration. Cross-sectional analysis of these images reveals the thickness of these layers to be approximately 4 nm.
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4. Discussion
Fig. 4. AFM phase image of SDS-treated inclusions showing 35-nm spherical structures on the surface of the ¢lter adjacent to the inclusions.
3.2. SDS studies Because our preliminary data suggested that the ‘rough’ inclusion possessed a boundary layer that is 4 nm thick (data from dried inclusions); and because this thickness agrees well with prior studies on the inclusion boundary layer[13,14], we pursued studies aimed at disruption of this boundary layer, and subsequent AFM imaging of this disruption. Inclusions treated with 0.5% (w/v) SDS at 45‡C for 5 min displayed a consistent deterioration process. First small holes were opened in the boundary layer, generally around the globular structures (data not shown). At this time, the parallel array could often be observed even though it was still underneath the boundary layer. As the deterioration progressed the boundary layer was made to look like a layer of Swiss cheese in that circular areas of deterioration could be imaged (Fig. 3). The holes gradually enlarged until there are only stringy strands of boundary layer left clinging to the surface of the inclusion. Strands of material were frequently seen connecting adjacent inclusions. Though the globular structures could be imaged early in the deterioration, they could not be imaged very late in the deterioration process. In images that contain inclusions that have largely lost their boundary layer, globular discs were observed scattered over the surface of the polycarbonate ¢lter near the inclusions (Fig. 4), but not in areas of the ¢lter that did not have inclusions. These structures were approximately the same diameter as the 35-nm globular structures imaged on intact inclusions.
The AFM data generated in this research project provides a di¡erent analysis of PHA inclusions from prior studies using electron microscopy. A large measure of this is due to the fact that the inclusions can be harvested and imaged with minimal preparation, thereby leaving structures intact that may be removed by the relatively harsh preparative procedures required in electron microscopy. Since AFM is a relatively new technology, there are issues that must be considered when applying the technology to a new system. For instance, AFM is a probe-limited technique that su¡ers from tip convolution e¡ects in which the geometry of the probe can distort the horizontal measurements made at the surface of an object [17^19]. This can be circumvented somewhat by using the distance between maximum height features to measure horizontal distances, but many horizontal distances must be viewed as estimates, rather than exact measurements. For convex samples there is an ‘edge’ e¡ect in which the probe does not maintain a consistent contact with the surface as it traverses down the side of the object and the angle of contact becomes increasingly non-perpendicular. For this reason, only the data gathered from the relatively £at center of the object was included in the analyses described in this report. Most of the images displayed in this report are the result of phase imaging. Phase imaging is a complementary scanning probe technique that can be correlated to di¡erences in hardness or elasticity of an object and has been widely used for polymeric materials [20]. The technology is subject to artifactual images and must be carefully compared with height images in order to be considered valid. Though phase images are largely used in this report (because of their clarity), each one was carefully compared against a representative height image before it was deemed to be valid. Our preliminary data allows us to build a somewhat speculative, but testable, model for inclusion structure and function. First, we suggest (based on our and others’ data) that it is proven at this point that there is a 3^4-nmthick boundary layer that surrounds every inclusion. Most likely the ‘rough’ inclusions imaged by AFM in our experiments have a boundary layer, whereas the ‘smooth’ inclusions have lost it. The chemical nature of this boundary layer is not known, but good arguments based on structure and the hydrophobicity of the interior of the inclusion have indicated that it is a lipid monolayer [13]. Given this, the next question that must be asked is how substrates gain access, and depolymerized products gain egress though this boundary layer. Our AFM images clearly show structures, reminiscent of porins (but with larger dimensions than known porins), which are possible centers for access/egress. This possibility is supported by the fact that we, and others, have isolated porin-like mol-
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ecules from the surface of inclusions [21,22]. The porinlike structures appear frequently and randomly on the inclusion and are connected by linear spokes that are about 4 nm in width. Beneath these structures can be seen parallel arrays that have a 7-nm spacing. It seems likely that the parallel arrays are the polymer itself that is being imaged. PHAs form a well-known lamellar spherulitic microstructure [23^25] and we have observed a structure similar to these parallel arrays in spun-cast thin ¢lms of PHBco-V [26]. We further suggest that globular structures traversing the surface of the parallel arrays are the protein machinery that mediates structure, synthesis, and depolymerization. As such, a large portion of this is likely to be composed of PhaP. The estimated width of PhaP, based on its molecular mass, agrees with the width of the linear structures (4 nm). These structures must not be very tightly bound to the polymer because inclusions that have lost their boundary layer also lose this network. One explanation for this phenomenon would be that the structures are linked to the boundary layer. The ease with which these structures are lost o¡ the inclusion surface would explain why they have not been imaged by electron microscopy with its inherently rigorous preparation procedure. Based on the commonality of membrane-spanning functional tunnel complexes (such as ATPase) in biology, we suggest that the spherical structures containing the central pore are synthesis/depolymerization centers. Therefore, in this model as the inclusion grows new protein networks are laid across the surface and these synthesis/ biosynthesis centers are added to these tracks by a process that is unknown. As sites of synthesis and degradation it would be expected that PHA synthetic and/or degradative enzymes are transiently or permanently associated with this complex. The globular structures that can be seen littering the surface of the ¢lter after SDS treatment are intriguing, but problematic. Other than the fact that they are of the same diameter as the porin-like structures seen on the surface of the inclusion, there is no data to suggest that this is their origin. However, we are experimentally pursuing these as possible protein complexes for synthesis and depolymerization, which are held in place by a protein sca¡olding system. This model agrees with the Mayer and Hoppert [13] model with regard to the lipid monolayer boundary layer, but di¡ers in that it is more advanced with respect to the protein constituents of the inclusion, which we postulate to be in an organized cytoskeletal-like structure. Furthermore, the Mayer model (or any other model) does not conceive the possibility that organized ‘porin-like’ structures for the ingress and egress may be present on the inclusion surface.
Acknowledgements This research was supported by grants from the National Science Foundation (#0113202 and # 0071717).
References [1] Anderson, A.J. and Dawes, E.A. (1990) Occurrence, metabolism, metabolic role, and industrial uses of bacterial polyhydroxyalkanoates. Microbiol. Rev. 54, 450^472. [2] Steinbu«chel, A., Hustede, E., Liebergesell, M., Pieper, U., Timm, A. and Valentin, H. (1993) Molecular basis for biosynthesis and accumulation of polyhydroxyalkanoic acids in bacteria. FEMS Microbiol. Rev. 10, 347^350. [3] Gerngross, T.U., Reilly, P., Stubbe, J., Sinskey, A.J. and Peoples, O.P. (1993) Immunocytochemical analysis of poly-beta-hydroxybutyrate (PHB) synthase in Alcaligenes eutrophus H16: localization of the synthase enzyme at the surface of PHB granules. J. Bacteriol. 175, 5289^5293. [4] Wieczorek, R., Pries, A., Steinbuchel, A. and Mayer, F. (1995) Analysis of a 24-kilodalton protein associated with the polyhydroxyalkanoic acid granules in Alcaligenes eutrophus. J. Bacteriol. 177, 2425^ 2435. [5] York, G.M., Stubbe, J. and Sinskey, A.J. (2002) The Ralstonia eutropha PhaR protein couples synthesis of the PhaP phasin to the presence of polyhydroxybutyrate in cells and promotes polyhydroxybutyrate production. J. Bacteriol. 184, 59^66. [6] Potter, M., Madkour, M.H., Mayer, F. and Steinbuchel, A. (2002) Regulation of phasin expression and polyhydroxyalkanoate (PHA) granule formation in Ralstonia eutropha H16. Microbiol. SGM 148, 2413^2426. [7] Lundgren, D.G., P¢ster, R.M. and Merrick, J.M. (1964) Structure of poly(L-hydroxybuturic acid) granules. J. Gen. Microbiol. 34, 441^ 446. [8] Steinbu«chel, A., Aerts, K., Babel, W., Follner, C., Liebergesell, M., Madkour, M.H., Mayer, F., Pieper-Furst, U., Pries, A. and Valentin, H.E. et al. (1995) Considerations on the structure and biochemistry of bacterial polyhydroxyalkanoic acid inclusions. Can. J. Microbiol. 41, 94^105. [9] Stuart, E.S., Lenz, R.W. and Fuller, R.C. (1995) The ordered macromolecular surface of polyester inclusion bodies in Pseudomonas oleovorans. Can. J. Microbiol. 41, 84^93. [10] Stuart, E.S., Tehrani, A., Valentin, H.E., Dennis, D., Lenz, R.W. and Fuller, R.C. (1998) Protein organization on the PHA inclusion cytoplasmic boundary. J. Biotechnol. 64, 137^144. [11] Fuller, R.C., O’Donnell, J.P., Saulnier, J., Redlinger, T.E., Foster, J. and Lenz, R.W. (1992) The supramolecular architecture of the polyhydroxyalkanoate inclusions in Pseudomonas oleovorans. FEMS Microbiol. Rev. 103, 279^288. [12] Fuller, R.C. (1999) Microbial inclusions with special reference to PHA inclusions and intracellular boundary envelopes. Int. J. Biol. Macromol. 25, 21^29. [13] Mayer, F. and Hoppert, M. (1997) Determination of the thickness of the boundary layer surrounding bacterial PHA inclusion bodies and implications for models describing the molecular architecture of this layer. J. Basic Microbiol. 37, 45^52. [14] Boatman, E.S. (1964) Observations on the ¢ne structure of spheroplasts of Rhodospirillum rubrum. J. Cell Biol. 20, 297^306. [15] Sudesh, K., Gan, Z.H., Maehara, A. and Doi, Y. (2002) Surface structure, morphology and stability of polyhydroxyalkanoate inclusions characterised by atomic force microscopy. Polym. Degrad. Stab. 77, 77^85. [16] Schlegel, H.G., Vonbartheld, R. and Gottschalk, G. (1961) Formation and utilization of poly-beta-hydroxybutyric acid by Knallgas bacteria (Hydrogenomonas). Nature 191, 463. [17] Budashov, I.A., Kurochkin, I.N., Denisov, A.K., Skripniuk, V.V. and Shabanov, T.A. (1997) The use of atomic-force microscopy for the analysis of microbiological objects. Zh. Mikrobiol. Epidemiol. Immunobiol. 6, 11^15. [18] Ahimou, F.O., Touhami, A. and Dufrene, Y.F. (2003) Real-time imaging of the surface topography of living yeast cells by atomic force microscopy. Yeast 20, 25^30.
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D. Dennis et al. / FEMS Microbiology Letters 226 (2003) 113^119 [19] Giocondi, M.C., Milhiet, P.E., Lesniewska, E. and Le Grimellec, C. (2003) From cell imaging to molecular manipulation : biological applications of atomic force microscopy. Med. Sci. 19, 92^ 99. [20] Berquand, A., Mazeran, P.E. and Laval, J.M. (2003) In£uence of volume and surface properties on phase contrast in tapping mode atomic force microscopy. Surf. Sci. 523, 125^130. [21] Matsumoto, K., Matsusaki, H., Taguchi, K., Seki, M. and Doi, Y. (2002) Isolation and characterization of polyhydroxyalkanoates inclusions and their associated proteins in Pseudomonas sp 61^3. Biomacromolecules 3, 787^792. [22] Dennis, D.E. (2002) Unpublished results. [23] Murase, T., Iwata, T. and Doi, Y. (2001) Atomic force microscopy
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investigation of poly[(R)-3-hydroxybutyrate] lamellar single crystals : Relationship between molecular weight and enzymatic degradation behavior. Macromol. Biosci. 1, 275^281. [24] Murase, T., Iwata, T. and Doi, Y. (2001) Direct observation of enzymatic degradation behavior of poly[(R)-3-hydroxybutyrate] lamellar single crystals by atomic force microscopy. Macromolecules 34, 5848^5853. [25] Rossini, C.J., Arceo, J.F., McCarney, E.R., Augustine, B.H., Dennis, D.E., Flythe, M.D. and Baron, S.F. (2001) AFM studies on poly-3hydroxybutyrate. In: Macromolecular Symposia Advances in Scanning Probe Microscopy of Polymers (Tsukruk, V.V. and Spencer, N.D., Eds.). Wiley-VCH, Weinheim, Germany. [26] McCarnery, E.R. and Augustine, B.H. (1999) Unpublished results.
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Preliminary analysis of polyhydroxyalkanoate inclusions using atomic force microscopy Douglas Dennis a; , Caroline Liebig b , Tara Holley b , Kara S. Thomas b , Amit Khosla b , Douglas Wilson b , Brian Augustine c a
Department of Life Sciences, Arizona State University West, P.O. Box 37100, Phoenix, AZ 85069-7100, USA b Department of Biology, James Madison University, Harrisonburg, VA, USA c Department of Chemistry, James Madison University, Harrisonburg, VA, USA Received 5 May 2003 ; received in revised form 8 July 2003 ; accepted 15 July 2003 First published online 26 August 2003
Abstract Atomic force microscopy analysis of polyhydroxyalkanoate (PHA) inclusions isolated from sonicated Ralstonia eutropha cells revealed that they exhibit two types of surface structure and shape; rough and ovoid, or smooth and spherical. Smooth inclusions possessed linear surface structures that were in parallel arrays with 7-nm spacing. Occasionally, cracks or fissures could be seen on the surface of the rough inclusions, which allowed a measurement of approximately 4 nm for the thickness of the boundary layer. When the rough inclusions were imaged at higher resolution, globular structures, 35 nm in diameter, having a central pore could be seen. These globular structures were connected by a network of 4-nm-wide linear structures. When the inclusions were treated with sodium lauryl sulfate, the boundary layer of the inclusion deteriorated in a manner that would be consistent with a lipid envelope. When the boundary layer was largely gone, 35nm globular disks could be imaged laying on the surface of the filter beside the inclusions. These data have facilitated the development of a preliminary model for PHA inclusion structure that is more advanced than previous models. 7 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords : Atomic force microscopy; Poly-3-hydroxyalkanoate; Inclusion
1. Introduction Bacterial polyhydroxyalkanoates (PHAs) are biodegradable polyesters that are found as intracellular inclusions that are generally between 200 and 500 nm in diameter [1,2]. At present, very little is known about their structure and biogenesis. In Ralstonia eutropha, three proteins have been shown to exist at their surface, PhaC, PhaP and PhaR. PhaC is the PHA synthase and is responsible for the formation of the polyester from 3-hydroxy fatty acylCoA molecules [2,3]. PhaP is thought to be a structural protein because it is found in very large quantities in PHA-accumulating cells and has profound e¡ects on the shape and number of PHA inclusions when it is deleted or
* Corresponding author. Tel. : +1 (602) 543 6934; Fax : +1 (602) 543 6073. E-mail address : [email protected] (D. Dennis).
overexpressed [4]. PhaR regulates the expression of PhaP [5,6]. Early electron microscopic studies (carbon replica) on inclusions from Bacillus megaterium supported the existence of a membrane layer at the surface of the inclusion [7], but later studies on inclusions from Pseudomonad species imaged a regular lattice-like surface architecture that is reminiscent of a bacterial S-layer [8^12]. Because of this, some researchers suggested that a protein lattice covers the inclusions, most logically comprised of PhaP. The possibility of whether the boundary layer is either membrane or protein has been somewhat clari¢ed in that researchers have determined that boundary layer thickness is 4 nm [13,14], a fact that would seem to exclude the possibility of a lipid bilayer surrounding the inclusion because lipid bilayers are approximately 8 nm in thickness. Mayer and Hoppert have o¡ered strong theoretical arguments based on hydrophobic interactions of membranes as to why this boundary layer is most probably a lipid monolayer. Nonetheless, de¢nitive evidence on the composition of the boundary layer and the arrangement of proteins at its sur-
0378-1097 / 03 / $22.00 7 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/S0378-1097(03)00610-4
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face is scarce, and several models of inclusion structure continue to exist [2,8,9,11,12]. Recently, atomic force microscopy (AFM) studies on inclusions from Comomonas acidovorans have given support to the existence of an envelope around the inclusion [15]. The possible existence of a lipid boundary layer is quite interesting because a corollary implication is that there would have to be structures that would allow for the ingress of substrates for polymerization, as well as egress of depolymerized monomers to be released into the cytoplasm. The recent AFM work also noted the presence of uncharacterized 30-nm ‘globular’ structures on the surface of the inclusions. The use of AFM to analyze PHA inclusions surfaces is particularly appropriate because of the minimal amount of preparation necessary to image the inclusions. Past inclusion models su¡er from the fact that they were developed from electron micrographs of PHA inclusions that required signi¢cant sample preparation and manipulation prior to microscopy, possibly disrupting the integrity of the inclusion surface. In this study, we present AFM images that con¢rm the existence of the boundary layer and which reveal a network of organized structures on the surface of the inclusion that may function in structure and metabolism, as well as facilitating movement of molecules through the boundary layer.
2. Materials and methods 2.1. Bacterial strains and culture conditions R. eutropha was grown in Luria broth containing no NaCl (LBN) for routine growth and was maintained on LBN agar. For induction of PHA accumulation the bacteria was grown in Schlegel’s nitrogen-limited minimal media containing 0.1% (w/v) ammonium sulfate and 0.5% fructose [16]. 2.2. Inclusion preparation R. eutropha cultures were harvested when inclusions were clearly visible intracellularly (light microscopy), which was usually about 24 h post-inoculation when the Schlegel’s nitrogen-limited media was used. Five microliters of culture was added to 5 ml of 50 mM Tris (pH 8.0)^ 1 mM EDTA (or 50 mM Tris (pH 8.0)^0.15 M NaCl) and the suspension was added to a Falcon 2059 polypropylene tube and sonicated using an Tekmar 400 watt sonicator with a mid-sized probe. Sonication was done in an ice bath for 5 min using 5-s-on and 7-s-o¡ cycles. To rid the sonicated suspension of intact cells, the tube was centrifuged at 1000Ug for 10 min at 4‡C. The top 4 ml of the supernate was removed to a new tube and placed on ice. For AFM studies, varying amounts of this suspension were added to 10 ml of 50 mM Tris (pH 8.0)^1 mM EDTA and this was
¢ltered through a 250-nm pore size polycarbonate ¢lter. After the initial 10 ml, the ¢lter was washed with 2U5 ml of TE, followed by 5 ml of water. The ¢lter was immediately removed, adhered to a metal puck (12 mm; Ted Pella, Inc) and was directly imaged by AFM. 2.3. Sodium dodecyl sulfate (SDS)-mediated disintegration of boundary layer This procedure was the same as above, except that the 4 ml of supernate removed after the 1000Ug centrifugation was now added to 36 ml of a prewarmed (45‡C) SDS solution such that the ¢nal SDS concentration was 0.5% (w/v). The suspension was gently mixed and incubated at 45‡C for 5 min. Varying amounts of the suspension were removed and ¢ltered through 250-nm pore size polycarbonate ¢lters, followed by two 5-ml washes with deionized water. Filters were processed as above. 2.4. AFM AFM was performed on a Digital Instruments Multimode instrument operating in tapping mode to acquire both height and phase images simultaneously. All imaging was performed in air at room temperature using silicon tapping mode probes manufactured by Olympus and purchased from Digital Instruments. Typical resonance frequencies for the probes were V250^300 kHz. Phase imaging was obtained by applying a relatively large tapping force (low setpoint) to the surface of the inclusion bodies. Typical scan speeds were 0.3^0.7 Hz.
3. Results In general, two types of inclusions (Fig. 1A,B.) were observed by AFM. The majority of inclusions tended to be ovoid in appearance and have a ‘rough’ or ‘bumpy’ surface. Occasionally, another type of inclusion that is rounder and had a smoother surface was seen. The ‘smooth’ inclusions had a di¡erent pro¢le in that they are more ‘mushroom-like’ and did not extend as high above the surface of the ¢lter (cross-section data not shown). Generally, their height pro¢le was about half of the pro¢le of rough inclusions. The proportion of smooth inclusions in the preparation increased if, 1) bacterial cells stored at 4‡C for several days were used, 2) the cells were subjected to longer periods of sonication, or 3) higher levels of EDTA were used (5 or 10 mM). When the ¢lters containing inclusions were stored at room temperature for a week and re-imaged, wide ¢ssures or cracks on the surface of the inclusion could be seen, presumably due to drying. The surface of the sublayer at the bottom of these ¢ssures is consistently 4 nm below the surface of the ‘rough’ inclusion (cross-section analysis, data now shown).
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Fig. 1. AFM images of PHA inclusions. AFM height (A) and phase (B) images of two types of PHA inclusions isolated from sonicated cells. In the height image the brightness of the image is directly related to the height of the inclusion. Terracing at the edge of the inclusion in the phase image is an artifact and should be disregarded. C: AFM high resolution phase image of the surface of a smooth inclusion showing 7-nm parallel arrays. D : AFM phase image of rough inclusion showing 35-nm globular structures with central pore.
3.1. High resolution studies When the ‘rough’ and ‘smooth’ inclusions were imaged using AFM at resolutions near the limit of the instrument, a di¡erent type of structure could be identi¢ed on the surface of the inclusion. The smooth inclusions had linear strands in parallel arrays traversing the surface of the inclusions (Fig. 1C). These parallel arrays did not extend uninterrupted across the surface of the inclusions, but could be seen traversing the surface of the inclusion in regions that were then interrupted by another set of linear arrays traversing a di¡erent direction. It would be analogous to the strokes of a paintbrush across the surface of the inclusion. The distance between the lines of the parallel
array was quite consistent and was measured by cross-sectional analysis at 7 nm (data not shown). The ‘rough’ inclusions exhibit quite a di¡erent surface morphology. The surfaces of these inclusions were populated with globular structures that were roughly spherical and which had a central pore (Fig. 1D). The globular structures were consistent in size, approximately 35 nm in diameter. Cross-sectional analyses of these structures revealed that the average pore diameter of the structure was about 15 nm (at the surface). The collar of the structure was about 12^15 nm in diameter and extended 4^12 nm above the surface of the inclusion (data not shown). Generally, many pores can be seen and they appear to be connected by linear structures that traverse the surface of the inclusion,
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Fig. 2. High resolution AFM phase image of PHA inclusion. Speci¢c structures mentioned in text: 1: Parallel arrays with 7-nm spacing; 2: linear network-like structures ; 3: porin-like structures with central pore. Network structures appear to be more prominent at the edge of scan because of tip convolution.
making it appear that there is a cytoskeletal structure encompassing the inclusion (Fig. 2). The average width of these structures is about 4 nm, but they can be wider, particularly at the points that they connect to the ‘porin-
like’ structures. In the highest resolution images, the same parallel linear arrays (7 nm spacing) seen on smooth inclusions could be imaged beneath the cytoskeletal-like network (Fig. 2).
Fig. 3. AFM phase images of inclusions treated with warmed 0.5% SDS. The left image shows the boundary layer in earlier stages of deterioration, where the right image shows an inclusion in the later stage of deterioration. Cross-sectional analysis of these images reveals the thickness of these layers to be approximately 4 nm.
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4. Discussion
Fig. 4. AFM phase image of SDS-treated inclusions showing 35-nm spherical structures on the surface of the ¢lter adjacent to the inclusions.
3.2. SDS studies Because our preliminary data suggested that the ‘rough’ inclusion possessed a boundary layer that is 4 nm thick (data from dried inclusions); and because this thickness agrees well with prior studies on the inclusion boundary layer[13,14], we pursued studies aimed at disruption of this boundary layer, and subsequent AFM imaging of this disruption. Inclusions treated with 0.5% (w/v) SDS at 45‡C for 5 min displayed a consistent deterioration process. First small holes were opened in the boundary layer, generally around the globular structures (data not shown). At this time, the parallel array could often be observed even though it was still underneath the boundary layer. As the deterioration progressed the boundary layer was made to look like a layer of Swiss cheese in that circular areas of deterioration could be imaged (Fig. 3). The holes gradually enlarged until there are only stringy strands of boundary layer left clinging to the surface of the inclusion. Strands of material were frequently seen connecting adjacent inclusions. Though the globular structures could be imaged early in the deterioration, they could not be imaged very late in the deterioration process. In images that contain inclusions that have largely lost their boundary layer, globular discs were observed scattered over the surface of the polycarbonate ¢lter near the inclusions (Fig. 4), but not in areas of the ¢lter that did not have inclusions. These structures were approximately the same diameter as the 35-nm globular structures imaged on intact inclusions.
The AFM data generated in this research project provides a di¡erent analysis of PHA inclusions from prior studies using electron microscopy. A large measure of this is due to the fact that the inclusions can be harvested and imaged with minimal preparation, thereby leaving structures intact that may be removed by the relatively harsh preparative procedures required in electron microscopy. Since AFM is a relatively new technology, there are issues that must be considered when applying the technology to a new system. For instance, AFM is a probe-limited technique that su¡ers from tip convolution e¡ects in which the geometry of the probe can distort the horizontal measurements made at the surface of an object [17^19]. This can be circumvented somewhat by using the distance between maximum height features to measure horizontal distances, but many horizontal distances must be viewed as estimates, rather than exact measurements. For convex samples there is an ‘edge’ e¡ect in which the probe does not maintain a consistent contact with the surface as it traverses down the side of the object and the angle of contact becomes increasingly non-perpendicular. For this reason, only the data gathered from the relatively £at center of the object was included in the analyses described in this report. Most of the images displayed in this report are the result of phase imaging. Phase imaging is a complementary scanning probe technique that can be correlated to di¡erences in hardness or elasticity of an object and has been widely used for polymeric materials [20]. The technology is subject to artifactual images and must be carefully compared with height images in order to be considered valid. Though phase images are largely used in this report (because of their clarity), each one was carefully compared against a representative height image before it was deemed to be valid. Our preliminary data allows us to build a somewhat speculative, but testable, model for inclusion structure and function. First, we suggest (based on our and others’ data) that it is proven at this point that there is a 3^4-nmthick boundary layer that surrounds every inclusion. Most likely the ‘rough’ inclusions imaged by AFM in our experiments have a boundary layer, whereas the ‘smooth’ inclusions have lost it. The chemical nature of this boundary layer is not known, but good arguments based on structure and the hydrophobicity of the interior of the inclusion have indicated that it is a lipid monolayer [13]. Given this, the next question that must be asked is how substrates gain access, and depolymerized products gain egress though this boundary layer. Our AFM images clearly show structures, reminiscent of porins (but with larger dimensions than known porins), which are possible centers for access/egress. This possibility is supported by the fact that we, and others, have isolated porin-like mol-
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ecules from the surface of inclusions [21,22]. The porinlike structures appear frequently and randomly on the inclusion and are connected by linear spokes that are about 4 nm in width. Beneath these structures can be seen parallel arrays that have a 7-nm spacing. It seems likely that the parallel arrays are the polymer itself that is being imaged. PHAs form a well-known lamellar spherulitic microstructure [23^25] and we have observed a structure similar to these parallel arrays in spun-cast thin ¢lms of PHBco-V [26]. We further suggest that globular structures traversing the surface of the parallel arrays are the protein machinery that mediates structure, synthesis, and depolymerization. As such, a large portion of this is likely to be composed of PhaP. The estimated width of PhaP, based on its molecular mass, agrees with the width of the linear structures (4 nm). These structures must not be very tightly bound to the polymer because inclusions that have lost their boundary layer also lose this network. One explanation for this phenomenon would be that the structures are linked to the boundary layer. The ease with which these structures are lost o¡ the inclusion surface would explain why they have not been imaged by electron microscopy with its inherently rigorous preparation procedure. Based on the commonality of membrane-spanning functional tunnel complexes (such as ATPase) in biology, we suggest that the spherical structures containing the central pore are synthesis/depolymerization centers. Therefore, in this model as the inclusion grows new protein networks are laid across the surface and these synthesis/ biosynthesis centers are added to these tracks by a process that is unknown. As sites of synthesis and degradation it would be expected that PHA synthetic and/or degradative enzymes are transiently or permanently associated with this complex. The globular structures that can be seen littering the surface of the ¢lter after SDS treatment are intriguing, but problematic. Other than the fact that they are of the same diameter as the porin-like structures seen on the surface of the inclusion, there is no data to suggest that this is their origin. However, we are experimentally pursuing these as possible protein complexes for synthesis and depolymerization, which are held in place by a protein sca¡olding system. This model agrees with the Mayer and Hoppert [13] model with regard to the lipid monolayer boundary layer, but di¡ers in that it is more advanced with respect to the protein constituents of the inclusion, which we postulate to be in an organized cytoskeletal-like structure. Furthermore, the Mayer model (or any other model) does not conceive the possibility that organized ‘porin-like’ structures for the ingress and egress may be present on the inclusion surface.
Acknowledgements This research was supported by grants from the National Science Foundation (#0113202 and # 0071717).
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