pubs.acs.org/Langmuir © 2009 American Chemical Society
Double-Shell Giant Vesicles Mimicking Gram-Negative Cell Wall Behavior during Dehydration Agnes Csiszar, Bernd Hoffmann, and Rudolf Merkel* :: :: Institute of Bio- and Nanosystems, Biomechanics (IBN-4), Research Centre Julich, 52425 Julich, Germany Received December 12, 2008. Revised Manuscript Received March 23, 2009 A biomimetic system modeling the behavior of Gram-negative bacteria under hyperosmotic stress was developed. To this end, we introduced a two-step electroswelling procedure for encapsulation of giant unilamellar vesicles by an additional membrane. Both membranes of the resulting double-shell vesicles (DSVs) were fluid. Additionally, the outer membrane was rigidified by a monolayer of streptavidin forming a two-dimensional crystal. For strong attachment of this protein layer, the outer membrane contained biotinylated lipids. This reinforced protein-lipid compound membrane served to model the assembly of the murein wall and outer membrane of Gram-negative bacteria. We characterized DSVs by confocal laser scanning microscopy. Furthermore, DSVs were exposed to hyperosmotic media (osmotic difference 0-1100 mosm/L), and the resulting shapes were analyzed. DSVs coated with streptavidin were much less deformed or destroyed by osmotic stress than bare DSVs or DSVs coated with noncrystalline avidin. Osmotically stressed DSVs coated with streptavidin displayed weak wrinkling of the outer membrane and formed small daughter vesicles of the inner membrane. Both features and the toughness against hyperosmotic stress are well described characteristics of Gram-negative bacteria.
Introduction Bacteria, like all living organisms, depend on water for their metabolism. As water readily passes through biomembranes, its flow into or out of the cell is controlled by the osmotic pressure difference between the outer medium and the cell cytoplasm. Optimal conditions for cell growth require an isoosmotic medium. In hyperosmotic media (outside osmotic pressure exceeds the internal one), excess water must be pumped out to reduce the pressure acting on the cell wall. In the opposite case (hypoosmotic media), cells must pump water against the osmotic pressure difference into the cell. In both cases, the cell is exposed to osmotic stress and tries to compensate by activating its osmotic protection mechanism.1,2 This mechanism and the compensation pressure range strongly depend on the membrane structure and membrane composition of the microorganism.3 For example, the osmotic protection of Gram-negative bacteria is based on their double cell wall structure. It protects the cell as follows: First, the plasma membrane senses the osmotic change of the environment and activates the regulatory system of cells to restore volume and turgor pressure.1,2 However, this cell reaction can balance only a limited change of osmotic pressure. In the case of Gram-negative cells, this limit is about 10 MPa,4 which corresponds to an osmolarity difference of approximately 4.4 osm/L. Osmotic pressure π is defined by an osmosis experiment where two solutions are separated by a semipermeable membrane. Only solutes that cannot pass the membrane contribute to the osmotic pressure, which is defined as the additional hydrostatic pressure that must be applied to the solution of higher osmolarity to stop water flow through the membrane. The osmolarity of a solution is
the sum of all concentrations of freely diffusing solutes that cannot pass the membrane. It is given in the unit osm/L. Osmolarity c and osmotic pressure π are connected by van t0 Hoff ’s law: π ¼ cRT 103
ð1Þ
where R is the gas constant and T is the absolute temperature. The additional factor of 1000 arises from the conversion of liters to m3. For osmotic pressures above the threshold of 10 MPa, a mechanical process is activated. The cell volume decreases while the cell membrane undergoes weak surface wrinkling. An additional increase of osmotic pressure causes the separation of the cytoplasmic membrane from the rigid outer membrane-murein wall complex, thereby producing the so-called plasmolysis space.5 Simultaneously, small endocytotic periplasmatic vesicles split from the cytoplasmic membrane to reduce its surface.6 At the end, an almost unwrinkled outer membrane and a relaxed cytoplasmic membrane with the plasmolysis space in between can be observed.7 In a pressure range of 10-40 MPa, the outer membrane surface becomes more wrinkled but no other basic change is observed. Osmotic pressures above 40 MPa induce a phase transition of lipid molecules from fluid crystalline phase to gel phase. This causes an immediate change of membrane fluidity. Simultaneously, cellular viability decreases drastically.4 From a biophysical point of view, cell response to mechanical stimuli is highly interesting. The mechanism of the mechanical protection of Gram-negative cells is still not yet elucidated. Furthermore, it is also unclear how important the mechanical protection against osmotic stress is for various types of microorganisms.3 Most intriguingly, mechanical protection could be based on active processes of the cell or on physicochemical principles without involving molecular pathways. Because the
*Corresponding author. (1) (2) (3) (4) 464.
Wood, J. Mol. Biol. Rev. 1999, 63, 230–262. Wood, J. Methods Enzymol. 2007, 428, 77–107. Mille, Y.; Beney, L.; Gervais, P. Biotechnol. Bioeng. 2005, 92, 478–484. Beney, L.; Mille, Y.; Gervais, P. Appl. Microbiol. Biotechnol. 2004, 65, 457–
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(5) Olijhoek, A.; Vaneden, C.; Trueba, F. J. J. Bacteriol. 1982, 152, 479–484. (6) Nobel, P. S. Biophysical Plant Physiology and Ecology; W H Freeman & Co: San Francisco, 1983. (7) Schwarz, H.; Koch, A. L. Microbiology (Reading, U.K.) 1995, 141, 3161– 3170.
Published on Web 4/23/2009
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separation of these pathways is difficult to realize in living cells, application of a model membrane system could solve these open questions. For example, a phospholipid vesicle system consisting of two separate lipid bilayers, one enclosed in the other, can mimic the double-shell construction of a Gram-negative cell wall as well as simulate the different mechanical properties of inner and outer membranes. To mimic the rigid outer membrane response to hyperosmotic pressures, Horton et al.8 used a model membrane system which contains biotin-functionalized giant unilamellar vesicles (GUVs) coated with a two-dimensional protein crystal form of streptavidin.9,10 Increasing osmotic pressure was generated by increasing glucose concentrations in the outer solution. These osmotically stressed streptavidin coated vesicles presented a crumpled shape, with regions of high membrane curvature, similar to a Gramnegative cell wall,7 but the shape deformation began at significantly lower hyperosmotic pressures. Another disadvantage of this model system is the absence of the cytoplasmic membrane and thereby the absence of possible membrane relaxation through membrane budding. The inner cytoplasmic membrane of bacteria was also mimicked using phospholipid vesicles. Its composition11 and packing, characteristics depending on temperature were studied, for example, by Lohner et al.12 Its behavior under thermal as well as osmotic stress was described by Wiese and co-workers.13,14 Their :: theory was detailed and experimentally supported by Kas and 15 16,17 18 and Pencer et al. These Sackmann, Seifert and co-workers, membrane systems focused on the properties of the cytoplasmic membrane in the absence of an outer membrane. In this work, we developed a model system to simultaneously mimic the rigid outer membrane, the highly elastic cytoplasmic membrane, and their mechanical behavior under hyperosmotic stress. The outer membrane was mimicked by a phospholipid bilayer labeled by a green fluorescent dye and also contained biotinylated lipids with negative charge. It was coated with crystalline streptavidin or, for comparison, with noncrystalline avidin. The inner membrane contained negatively charged lipids, neutral lipids, and red fluorescently labeled phospholipids. Thus, a twoshell membrane system was created as an approximation of a Gram-negative cell wall to mimic its behavior from a mechanical point of view. The shape of double-shell giant vesicles was observed by fluorescence laser scanning microscopy under isoosmotic conditions as well as under hyperosmotic stress. The changes in membrane curvature and overall shape were analyzed and compared with changes of pure unilamellar phospholipid vesicles and streptavidin tethered unilamellar vesicles. The influence of rigidity was explored by replacing the rigid crystalline streptavidin layer by a noncrystalline layer of avidin. Our results on double-shell vesicles (DSVs) with a crystalline outer layer closely resembled published observations of Gram-negative cell walls.7 (8) Horton, M. R.; Manley, S.; Arevalo, S. R.; Lobkowsky, A. E.; Gast, A. P. J. Phys. Chem. B 2007, 111, 880–885. (9) Ratanabanangkoon, P.; Gropper, M.; Merkel, R.; Sackmann, E.; Gast, A. P. Langmuir 2002, 18, 4270–4276. (10) Ratanabanangkoon, P.; Gropper, M.; Merkel, R.; Sackmann, E.; Gast, A. P. Langmuir 2003, 19, 1054–1062. (11) Gennis, R. B. In Biomembranes; Gennis, R. B., Ed.; Springer-Verlag: New York, 1989; p 21. (12) Lohner, K.; Latal, A.; Degovics, G.; Garidel, P. Chem. Phys. Lipids 2001, 111, 177–192. (13) Wiese, W.; Helfrich, W. J. Phys.: Condens. Matter 1990, 2, SA329–SA332. (14) Wiese, W.; Harbich, W.; Helfrich, W. J. Phys.: Condens. Matter 1992, 4, 1647–1657. :: (15) Kas, J.; Sackmann, E. Biophys. J. 1991, 60, 825–844. :: (16) Miao, L.; Seifert, U.; Wortis, M.; Dobereiner, H. G. Phys. Rev. E (A) 1994, 49, 5389–5407. (17) Seifert, U.; Berndl, K.; Lipowsky, R. Phys. Rev. A 1991, 44, 1182–1202. (18) Pencer, J.; White, G. F.; Hallett, F. R. Biophys. J. 2001, 81, 2716–2728.
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Materials and Methods Chemicals. To mimic mechanical properties and composition of cytoplasmic membranes of Gram-negative bacteria, 1,2dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1,2-dioleoylsn-glycero-3-phosphoethanolamine (DOPE) were chosen as main components. 1,2-dioleoyl-sn-glycero-3-(phospho-L-serine) (DOPS) yielded slightly negative charge of the inner vesicles, while 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (LissRhod-PE) was added to the mixture to visualize the model membranes in fluorescence microscopy. Lipid components were dissolved in chloroform and mixed to 1 mg/mL total lipid concentration in a molar ratio of DOPC/ DOPE/DOPS/LissRhod-PE = 1000/100/5/1. To produce a model for the outer bacterial membrane, a mechanically more rigid lipid, 1-stearoyl-2-oleoyl-sn-glycero-3-phosphocholine (SOPC) was chosen. Addition of negatively charged 1,2-dipalmitoyl-snglycero-3-phosphoethanolamine-N-(cap biotinyl) (capBioDPPE) enabled coupling of streptavidin to drastically increase the membrane rigidity. Green fluorescent tracer lipid N-(4,4-difluoro-5,7dimethyl-4-bora-3a,4a-diaza-s-indacene-3-propionyl)-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (Bodipy FL-DHPE) was also added to the lipid mixture. The lipids were dissolved and mixed in chloroform to 1 mg/mL total lipid concentration. The molar ratio used was DOPE/capBioDPPE/Bodipy FL-DHPE = 1000/100/1. All lipids were purchased from Avanti Polar Lipids Inc. (Alabaster, AL) in a purity of at least 99%, except Bodipy FL-DHPE, which was ordered from Invitrogen (Karlsruhe, Germany). To coat the vesicles by protein layers, streptavidin, streptavidin-Alexa Fluor 488 (streptavidin-Alexa488), avidin, and avidin-Alexa Fluor 488 were purchased from Invitrogen. All other chemicals such as sucrose, glucose, sodium carbonate, sodium bicarbonate, 2-(N-morpholino)ethanesulfonic acid (MES) buffer, and bovine serum albumin (BSA) were ordered from VWR (Darmstadt, Germany). All water used was deionized, ultrapure water produced by a Milli-Q Gradient A10 apparatus (Millipore, Bedford, MA). Vesicle Preparation: Two-Step Electroswelling. First, slightly negatively charged giant unilamellar vesicles were prepared by the electroswelling technique.19 A total of 10 μL of the lipid mixture DOPC/DOPE/DOPS/LissRhod-PE (1000/100/5/1) was carefully deposited on indium tin oxide (ITO) coated glass slides (Praezisionsglas & Optik GmbH, Iserlohn, Germany). The lipid films were then dried under vacuum for at least 1 h. For the first electroswelling step, the plates were placed in a chamber containing 2 mL of 300 mM sucrose solution (300 mosm/L) in pure water and separated by a 1 mm Teflon spacer. To exclude a possible effect of pH gradients across the vesicle membranes applied in the subsequently preparation steps, in some experiments, the pH of sucrose solution was adjusted to pH 5.5 by 0.5 M MES buffer or to pH 10 by 20 mM sodium carbonate/sodium bicarbonate buffer. The ionic strength of the swelling solutions remained below 10 mM. Vesicles were swollen in an alternating electric field of 1.5 V at 10 Hz for 2 h. After the first swelling step, ITO plates were replaced by two horizontally arranged ITO plates coated with a strongly negatively charged lipid mixture of SOPC/ capBioDPPE/Bodipy FL-DHPE (1000/100/1). Plates were incubated for 30 min in the vesicle solution that was formed in the previous electroswelling step to allow contact between lipid layers on the glass slides and preformed GUVs by adhesion. To increase the amount of GUVs close to the slide, incubation was performed in the presence of isoosmolar glucose solution which exhibits a lower density than the sucrose filled vesicles. Subsequently, electroswelling was repeated with the same parameters as before. To observe vesicles, 50 μL of the suspension was transferred to 2 mL of 300 mM glucose solution (300 mosm/L). The osmolalities (19) Angelova, M. I.; Soleau, S.; Meleard, Ph.; Faucon, F.; Bothorel, P. Prog. Colloid Polym. Sci. 1992, 89, 127–131.
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of glucose and vesicle solutions were determined using an Osmomat 030 freezing point depression osmometer by Gonotec (Berlin, Germany) and converted to osmolarities assuming a density of water of 1 kg/L. Proof of Double-Shell Vesicle Formation. To prove the existence of double-shell vesicles, control experiments were performed. For this purpose, both lipid mixtures were prepared without fluorescent dyes. After 2 h swelling of slightly negatively charged GUVs (DOPC/DOPE/DOPS = 1000/100/5) in 300 mM sucrose solution, Alexa488 fluorescent dye was added to the vesicle suspension at a high concentration of 1 μg/mL. After dye mixing with the suspension, vesicles were adhered to ITO slides covered by the strongly negatively charged lipid mixture (SOPC/ capBioDPPE = 1000/100). After 30 min of incubation, the second swelling step was started. To observe vesicles, 2 μL of the suspension was transferred to 2 mL of 300 mM glucose solution. Furthermore, to test for interaction of lipid mixtures and Alexa488, the two types of vesicles were swollen alone and incubated in a solution of 1 μg/mL Alexa488 also for several hours. The samples were investigated under the same conditions as above. Vesicle Coating with Proteins. For most experiments, the vesicles were coated with streptavidin. Before mixing of vesicles and protein, the pH value of protein solution was adjusted to pH 5.5 by 0.2 M MES/HCl buffer with an osmolarity equal to the vesicle solution. Vesicles were incubated in 70 μg/mL protein solution for at least 3 h. For some control experiments, vesicles were coated with Alexa488 labeled streptavidin (streptavidin/ streptavidin-Alexa488 = 1:1) at pH 5.5. In another case, vesicles were coated with avidin the same way as described above, but MES buffer was replaced by 20 mM sodium carbonate/sodium bicarbonate buffer at pH 10. The fluorescence signal was analyzed by microscopy. Imaging and Data Analysis. The observation chamber was coated with BSA (1 mg/mL) for 10 min and washed subsequently. The coated chamber was filled with 2 mL of 300, 650, 1000, or 1400 mosm/L glucose solution and 50 μL of the vesicle suspension (300 mosm/L), producing approximately 0, 350, 700, and 1100 mosm/L hyperosmotic pressures. Vesicles were imaged immediately after osmotic shock application for up to 1 min to 2 h using a laser scanning microscope LSM 710 (Carl Zeiss MicroImaging GmbH, Jena, Germany) equipped with an argon ion laser (488 nm) and a helium-neon laser (543 nm). To detect the fluorescent signal of LissRhod (excitation 543 nm), a long pass filter LP600 nm was used, and for Bodipy FL (excitation 488 nm) a band-pass filter BP500-520 nm was used. The microscope was focused on the vesicles using a 40/1.20 C-Apochromat water immersion objective (Carl Zeiss MicroImaging GmbH). Images were recorded by sequential imaging of the two detection channels with a resolution of 512 512 pixels. Pixel size was 154 nm. To record the statistical size distribution and occurrence of the three types of vesicles, the same region of the sample was recorded with two different focus planes, one for vesicles with a diameter smaller than 10 μm and another for the larger population. The overlay picture of the two detector channels was analyzed and edited with the LSM 710 ZEN software by Carl Zeiss MicroImaging GmbH. Additionally, Origin 7G (OriginLab Co., Northampton, MA) was used for statistical evaluations.
Results Vesicle Preparation. Formation of giant vesicles composed of two independent bilayers is a major tool to mimic the mechanics of Gram-negative cell walls. For the development of such a system, negatively charged GUVs labeled with LissRhod were swollen in the first preparation step. After preparation, the vesicles were deposited by gravity onto horizontally arranged ITO slides covered with also negatively charged, Bodipy FL labeled lipids. During the second swelling, the electric field accelerated the Langmuir 2009, 25(10), 5753–5761
Figure 1. Fluorescence micrographs of all three types of vesicles formed by two-step electroswelling. The same location within a sample was imaged in all three cases. (A) unilamellar vesicle labeled with LissRhod (red), (B) unilamellar vesicle labeled with Bodipy FL (green), and (C) double-shell giant vesicle (red and green simultaneously). Here, most likely the green fluorescent signal is weakened by energy transfer from Bodipy FL to LissRhod. The images represent quite well the size distribution and frequency of the three types of vesicles at the end of the swelling process. Scale bars = 10 μm.
detachment of vesicles. Green fluorescent unilamellar vesicles as well as preformed red fluorescent vesicles encapsulated within green fluorescent lipid bilayers were formed. At the end of the two-step electroswelling process, three different types of vesicles were observed by fluorescence microscopy: slightly negatively charged unilamellar vesicles (GUVs) identified by their red fluorescent signal (Figure 1A), strongly negatively charged unilamellar vesicles (GUVs) displaying green fluorescence (Figure 1B), and so-called double-shell vesicles (DSVs) emitting red and green fluorescent signals simultaneously (Figure 1C). DOI: 10.1021/la8041023
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Figure 2. Three types of vesicles formed by two-step electroswelling. Here, the second swelling step was performed in dye solution. (A) negatively charged vesicle filled with 300 mosm/L sucrose solution, (B) strongly negatively charged vesicle filled with 300 mosm/L sucrose solution and Alexa488 at 1 μg/mL concentration, and (C) double-shell vesicle with a water layer containing Alexa488 between the two shells. To reduce background intensity, the sample was diluted 1:1000. Scale bars = 10 μm.
Compared to the signals of the unilamellar vesicles in both the LissRhod and Bodipy FL detector channels, the signal of these double-shell vesicles appeared weaker for Bodipy FL and similarly intense for LissRhod. Proof of Double-Shell Vesicle Formation. The existence of double-shell vesicles was proven by an additional experiment in which the lipid bilayers remained unlabeled but the second swelling solution contained green fluorescent dye (Alexa488) in high concentration to label the water layer enclosed between the two lipid bilayers. As result, the first prepared negatively charged vesicles always remained unlabeled (Figure 2A). In contrast, the strongly negatively charged vesicles were swollen during the second preparation step and therefore enclosed a highly fluorescent solution (Figure 2B). In double-shell vesicles, the preformed vesicles were encapsulated in strongly negatively charged lipid bilayers with a highly fluorescent water layer in between (Figure 2C). To image all three types of formed vesicles, a dilution of 1:1000 was used. At this enhanced signal to background ratio, the encapsulated fluorescently labeled water layer was clearly visible. In contrast, at this high dilution of background fluorescence, the imaging of preformed vesicles was barely possible. To exclude the possibility of interaction between charged lipid membranes and Alexa488, unilamellar vesicles were prepared from both lipid mixtures and incubated with Alexa488 for several hours. These experiments did not show any measurable enrichment of the dye at the membrane surfaces (data not shown). Size Distribution and Occurrence of the Three Types of Vesicles. The frequency of occurrence and size distribution of all three vesicle types was determined by fluorescence microscopy. To distinguish vesicles with diameters above and below 10 μm, two images with different focus positions were taken for all regions of the sample. For the analysis, only diameters of vesicles that were in focus were measured to precisely determine the size distribution and simultaneously to avoid double counting of vesicles. Analysis after the first swelling step showed a Gaussian size distribution of GUVs (A) in the LissRhod channel (red) with an average diameter of 16.6 ( 6.9 μm and the absence of vesicles in the green Bodipy FL channel (Figure 3I and Table 1). After the second swelling step, the average diameter of vesicles detected in the LissRhod channel (red) was decreased to 11.5 ( 4.9 μm (Figure 3II) while GUVs in the Bodipy FL channel (green) appeared more or less in the same number as the red GUVs with an average diameter of 16.4 ( 8.1 μm. Their size showed a Gaussian distribution as well (Figure 3II). The third type of vesicle observed was the double-shell type (DSV), which appeared in the red and green detector channels simultaneously, resulting in an orange signal in the overlay. Its average diameter was found to 5756
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Figure 3. (I) Size distribution of unilamellar vesicles (GUV A) after the first swelling step under isoosmotic pressure. (II) Size distribution of unilamellar vesicles (GUV (A) and GUV (B)) as well as of double-shell vesicles (DSV (C)) after the second swelling step under isoosmotic conditions in sucrose solution and (III) in additional glucose solution. Vesicle types (A, B, and C) are defined in Figure 1. The size distribution of each vesicle population was fitted by a Gaussian function. The lines correspond to these Gaussian fits.
be 19.1 ( 5.9 μm, slightly higher than that of unilamellar vesicles, but its amount was only ∼1/7 of the entire vesicle population (Table 1). The general features of vesicle size distribution during a two-step swelling process were reproducible, while the absolute values strongly scattered depending on the swelling parameters. To increase the amount of DSVs, preformed GUVs were incubated in glucose solution on the ITO plates before the second swelling process. The resulting DSVs had a larger diameter of 25.3 ( 15.2 μm than with sucrose during the second swelling step, and their fraction increased related to the GUVs. A Gaussian fit of their distribution showed an asymmetrical tail in the higher diameter range, that was also typical for adhered GUVs (Figure 3III and Table 1). This adhesion reduced most likely the formation of free GUVs during the second swelling step and diminished their average diameter to 13.2 ( 5.1 μm. Langmuir 2009, 25(10), 5753–5761
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Table 1. Statistical and Gaussian Fit Parameters of the Three Vesicle Populations after the First and Second Electroswelling Processa vesicle type
no.
Dves ( sd ( μm)
DGauss ( μm)
1. swelling
GUV (A)
155
16.6 ( 6.9
15.5
1
2. swelling
GUV (A) GUV (B) GUV (C)
151 159 55
11.5 ( 4.9 16.4 ( 5.9 19.1 ( 5.9
10.7 14.8 20.1
0.39 0.44 0.16
AGauss
GUV (A) 150 17.2 ( 11.4 13.0 0.34 GUV (B) 155 13.2 ( 5.1 12.5 0.44 GUV (C) 65 25.3 ( 15.2 21.2 0.22 a Vesicle types (A, B, and C) are defined in Figure 1. Dves and sd are mean vesicle diameter and standard deviation as calculated from the measured distributions. The size distribution of each vesicle population was also fitted by a Gaussian function. The resulting fit parameters for diameters (DGauss) and relative occurrence of the respective vesicle type (AGauss) are given. 2. swelling in glucose solution
Figure 4. Effect of hyperosmotic pressure on GUVs labeled with LissRhod (red), streptavidin tethered GUVs labeled with Bodipy FL (green), and double-shell giant vesicles decorated with streptavidin (red and green overlapping signals (orange)). The inner vesicles were labeled by LissRhod, and the outer membrane with Bodipy FL. The hyperosmotic pressure differences were 0, 350, 700, and 1100 mosm/L. All images represent different vesicles. Scale bars = 5 μm.
Vesicle Morphology under Isoosmotic Conditions. In order to mimic the mechanics of stiff outer membranes of Gram-negative bacteria, vesicles were coated with streptavidin after the second swelling step. Since green fluorescence labeled and double-shell vesicles with green and red fluorescence always contained biotinylated lipids at a molar ratio of 0.1 at their surface, formation of biotin-streptavidin bonds was possible. The coating process was furthermore controlled by swelling nonfluorescent vesicles and coating them with Alexa488 labeled streptavidin. Confocal fluorescence microscopic analyses revealed a complete protein layer on the surface of these vesicles (data not shown). In contrast, red fluorescent vesicles remained uncoated due to their lack of biotinylated lipids. In the absence of streptavidin, thermally induced membrane fluctuations were observed, which slightly deformed the overall spherical shape of these vesicles (Figure 4, GUV at 0 mosm/L). On streptavidin tethered vesicles, no membrane fluctuations were observed, indicating more rigid and stiff structures compared to the pure phospholipid membranes (Figure 4, STR+GUV and STR +DSV at 0 mosm/L). Vesicle Morphology under Hyperosmotic Conditions. Pure phospholipid vesicles without streptavidin on their surface strongly fluctuated even at low hyperosmotic pressure (up to 2-3 mosm/L), and many small daughter vesicles were observed. Inside Langmuir 2009, 25(10), 5753–5761
Figure 5. Double-shell giant vesicle at 1100 mosm/L hyperosmotic pressure. The inner vesicle was labeled by LissRhod, and the outer membrane by Bodipy FL. Double-shell vesicles (DSVs) without surface proteins underwent an outside budding process under hyperosmotic conditions. The two membranes stuck together in newly formed nanotubes (yellow signal), or they formed separately new buds and tubes (green and red signals). An avidin coating on the double-shell vesicle surface (AV+DSV) could not inhibit the budding of the outer membrane (green signal) together with the inner membrane (red signal) during osmotic stress. In contrast, a vesicle coated with a crystalline streptavidin layer on the outer membrane surface (STR+DSV) showed a slight asymmetrical shape deformation without membrane budding (green signal), while the inner membrane released the osmotic pressure, separately forming daughter vesicles (red signal). Scale bars = 5 μm.
budding processes yielded small vesicles, which were observed inside the mother vesicles (for example, Figure 4, GUV 350 and 700 mosm/L). At the same time, small vesicles were formed and split up from the mother vesicles during outside budding processes (for example, Figure 4, GUV 1100 mosm/L), drastically decreasing the average vesicle diameter. Both processes occurred simultaneously at the used hyperosmotic pressures. However, the frequency of outside budding increased with increasing hyperosmotic pressures. Double-shell vesicles without a protein layer on the surface underwent budding under hyperosmotic pressures as well (Figure 5, DSV). In most cases, the two layers moved outside the core vesicle, like hundreds of tentacles or filaments as a DOI: 10.1021/la8041023
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Csiszár et al. Table 2. Relative Occurrence of Streptavidin Coated DSVs at Hyperosmotic Pressures of 0, 350, 700, and 1100 mosm/La fraction of DSVs (N = 49) ( sd at a hyperosmotic pressure of
shape of DSVs
0 mosm/L
350 mosm/L
750 mosm/L
1100 mosm/L
spherical 1 ( 0.00 0.81 ( 0.10 0.22 ( 0.10 0.10 ( 0.10 spherical with budds 0 0.16 ( 0.08 0.67 ( 0.22 0.18 ( 0.11 deformed with budds 0 0.03 ( 0.03 0.04 ( 0.03 0.22 ( 0.05 collapsed 0 0 0.07 ( 0.31 0.50 ( 0.09 a Results were taken from three independent preparations. Four different morphologies were defined: spherical shape, spherical shape with simultaneous budding of the inner layer, deformed shape with budding of the inner layer, and collapsed outer layer, with budding of the inner layer.
response to the large surface excess. In some cases, a simultaneous budding of the two lipid bilayers was also observed. Unilamellar vesicles with streptavidin on the surface (Figure 4, STR+GUV) exhibited a uniformly distributed pattern of folds for osmotic pressures of 350 mosm/L, but their overall shapes remained approximately spherical. Above a hyperosmotic pressure of 350 mosm/L, membrane folding was followed by a strong shape deformation and the vesicles became more asymmetric. In the high pressure range (up to 700 mosm/L), no further fundamental shape changes were observed. However, already existing wrinkles became deeper. Total collapse of these membranes was not observed in the analyzed pressure range. Streptavidin coated GUVs filled with Alexa488 solution displayed wrinkling but did not leak at high pressure with an inner solution that remained completely enclosed in the collapsed membrane capsule. The observed morphological changes of streptavidin crystal layers were exclusively caused by osmotic stress, and they were mainly irreversible. Interestingly, most double-shell vesicles appeared unchanged under hyperosmotic pressures lower than 350 mosm/L with neither shape deformations of the streptavidin layer, like unilamellar vesicles tethered by streptavidin, nor membrane budding, as found for pure phospholipid vesicles (Figure 4, STR+DSV). Moreover, the absence of membrane fluctuation as well as additional control experiments using fluorescently labeled streptavidin showed a successful surface coating. The first statistically significant morphological changes of streptavidin coated doubleshell vesicles appeared around 700 mosm/L hyperosmotic pressures. Here, the inner phospholipid membranes formed small daughter vesicles inside the mother vesicles. Most of them were firmly attached to the main membrane surface, but some vesicles at least partially detached into the vesicle lumen. In these cases, it could not be exactly determined if the detachment was complete or whether a fine lipid nanotube connected the daughter vesicles and the mother membrane. Increasing the hyperosmotic pressures to 1100 mosm/L, first surface wrinkling and asymmetrical shapes of the outer membrane accompained by budding of the inner membrane appeared. Furthermore, the complete leakage of the outer streptavidin layer upon streptavidin capsule collapse was observed and the inner elastic membrane budded out from the capsule, keeping the vesicle’s lumen intact (Table 2 and Figure 4, STR+DSV). Similar surface wrinkling had already been observed for unilamellar vesicles coated by streptavidin at about 100 times lower hyperosmotic pressures. Upon changing the surface coating from a crystalline to a noncrystalline protein layer, here avidin, similar but not the same membrane behavior as described above was observed. The inner membrane released pressure forming small vesicles into the vesicle lumen, but most of the daughter vesicles were strongly attached to the main membrane surface. Excess outer membrane coated with avidin in most cases accumulated in small lumps on the vesicle surface at the same spots where daughter vesicles were formed. 5758
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Figure 6. Streptavidin coated unilamellar and double-shell giant vesicles at 1100 mosm/L hyperosmotic pressure. Alexa488 containing buffer solution filled the unilamellar vesicle (top) and the gap between inner and outer membranes in the DSV (bottom). The unilamellar vesicle shrank strongly but completely retained its dye content (STR+GUV), while the double-shell vesicle with a water layer containing Alexa488 between the two layers just moderately shrank in comparison to the unilamellar vesicle. The two shells split up, causing an increased intermembrane volume. Note that the inner membrane strongly budded while the outer membrane was just slightly deformed.
In a hyperosmotic pressure range from 350 to 1100 mosm/L, some of them formed also small vesicles or nanotubes sticking to the inner membrane (Figure 5, AV+DSV). Similar to the morphological changes of streptavidin coated DSVs, these changes remained irreversible upon back transfer into isoosmotic solution (not shown). To test the integrity of membranes, GUVs and DSVs were prepared without fluorescent labeling of the membrane but with additional fluorescent dye dissolved in the second swelling solution. These vesicles were exposed to the same hyperosmotic pressures as described above. In the case of streptavidin coated GUVs, the same surface wrinkling and shape deformation processes were observed as without dye content. Even under high hyperosmotic pressure (1100 mosm/L), no leakage of interior dye solution was detected, indicating the absence of permanent pores (Figure 6, STR+GUV). DSVs coated by streptavidin showed the same permeation behavior but different shape transformations as streptavidin coated GUVs. Up to 700 mosm/L hyperosmotic pressure, deformation of streptavidin capsules with inner membrane budding was typical. Here, a separation of the inner and outer bilayers was observed with an increasing gap labeled with fluorescent dye. The inner membrane budding was exclusively detected in the bright field channel of the microscope (Figure 6, STR+DSV). The dye solution was completely enclosed between the two membranes, and no fluorescence signal could be detected in the bulk. Langmuir 2009, 25(10), 5753–5761
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Discussion A new model membrane system has been developed to mimic Gram-negative cell wall behavior during dehydration. It consists of two unilamellar phospholipid vesicles, one encapsulating the other. Their membranes are separated by repulsive electrostatic forces. The inner vesicle mimics the flexible inner cytoplasmic membrane of cells. The outer vesicle serves as a model for both the rigid outer cell membrane and the rigid murein wall, simultaneously. For this it is modified by a two-dimensional crystalline surface layer of streptavidin. To manufacture such double-shell vesicles, two lipid bilayers have to be brought close enough to enable the encapsulation but must be kept apart to inhibit membrane fusion between the two layers. For this reason, a new preparation process had to be developed with preformed giant unilamellar vesicles deposited on a layer of phospholipids of identical charge. The two membranes are pushed together by gravity and kept separated by repulsive electrostatic forces. The two lipid bilayers are separated by a thin water layer, whose presence was proven by fluorescent labeling of the solution. The fluorescent dye was not noticeably enriched at the membrane surfaces as illustrated in Figure 2A. The water layer thickness was not exactly determined, here but the distance between two neutral lipid bilayers in multilamellar vesicles is about 10-40 A˚ depending on the membrane composition.20,21 Presumably, the water layer thickness between similarly charged bilayers, like in our case, is further increased by the repulsive forces acting between the layers. Fluorescent labeling of both lipid layers with Bodipy FL and LissRhod yielded fluorescence energy transfer between the two bilayer leaflets which were separated by a thin water layer. Such energy transfer can occur without photon emission by fluorescence resonance energy transfer (FRET) if the distance of the two dyes is less than 60 A˚,22 or with photon emission and reabsorption. Because an exact elucidation of the energy transfer mechanism was not necessary in order to use the developed model system for cell membrane mimicking, no further investigations were carried out. In both transfer mechanisms, the excitation energy is transferred from Bodipy FL to LissRhod molecules, resulting in substantially weaker signal intensity in the Bodipy FL channel and a slightly increased intensity in the LissRhod channel as compared to unilamellar vesicles labeled only by Bodipy FL or LissRhod (Figure 1C). During the second electroswelling step to form double-shell giant vesicles, there were always some preformed unilamellar vesicles which did not adhere to the lipid layers on the glass surface. Moreover, lipid layers were only partially covered by adherent GUVs. In both cases, unilamellar vesicles were formed. Depending on the fluorescent dye incorporated in the membrane, their signals appeared in only one detector channel (Figure 1A and B). The vesicle size distributions after the two-step swelling process clearly showed a decrease of the average diameter of preformed unilamellar vesicles (Table 1 and Figure 3I and II). This shift was eliminated by incubation of the preformed vesicles in an isoosmolar glucose solution on the ITO plates before the second swelling step (Table 1 and Figure 3I and III). The complete adhesion of preformed GUVs strongly influenced the GUV population formed during the second swelling process. The (20) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P.; Parsegian, V. A. Biophys. J. 1982, 37, 657–665. (21) Cevc, G. Phospholipid Handbook; Marcel Dekker, Inc.: New York, 1993. (22) Yang, J.; Chen, H.; Vlahov, I. R.; Cheng, J. X.; Low, P. S. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 13872–13877.
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reduced uncovered surface resulted in vesicle formation only with significant decreased size and narrower size distribution. Interestingly, the induced adhesion of preformed GUVs did not significantly influence the population of DSVs. The interplay of the two opposing forces, gravity and electrostatic, during DSV swelling could not be fully clarified here. Even though the formation of DSVs without sufficient incubation time on the second lipid layer on ITO plates was not observed, an additional increase of gravity using glucose as second swelling solution neither induced significantly more DSVs nor changed vesicle population distribution (Table 1 and Figure 3). On the other hand, the repulsive forces between the two lipid bilayers appear quite high for the necessary close contact, but experiments with negatively and positively charged layers (data not shown) resulted in just one mixed lipid layer instead of two layers, one encapsulating the other. Thus, a certain amount of intermembrane repulsion seems necessary for DSV formation. In addition to the forces acting during swelling, the membrane composition can also play an important role in DSV formation. Here, it was important to find balance between molecules which brought the two leaflets in close proximity but did not induce their fusion. Presumably, the bulky headgroup of the biotinylated DPPE (capBioDPPE) was instrumental to prevent fusion. Although a double-shell membrane structure is essential to mimic Gram-negative cell membrane behavior under dehydration, the mechanical properties of the outer membrane are not well modeled yet. The typical outer membrane of Gram-negative cells is stiffened by a dense layer of surface proteins and serves as an adequate barrier against mechanical influences of the environment. To model the behavior of this outer cell membrane under hyperosmotic stress, the outer bilayers of our double-shell vesicles were coated by streptavidin. Before turning to the influence of streptavidin coating, the properties of uncoated vesicles under hyperosmotic stress are discussed. Under hyperosmotic conditions, the outer osmotic pressure was higher than the osmotic pressure inside the cell, or in our case inside the vesicles. This pressure difference induced a water flow from inside to outside reducing the enclosed volume, whereas the membrane surface remained constant. The driving force of osmosis was the osmotic pressure difference between the two sides. The changed volume to surface ratio induced bending and shear stress in the membrane.13,14 The membrane relaxation process depended on mechanical properties. For example, a highly elastic and flexible membrane, such as the cytoplasmic membrane of Gram-negative cells or the pure phospholipid membrane in our model system, was able to reduce the compression energy by forming small daughter vesicles or buds. The process is well-known in biology6 and is called membrane blebbing. However, the word budding is also frequently used by authors from the field of vesicles and membranes.15-17,23 Therefore, we use the term budding as well. Budding is highly favored by membranes containing different types of lipids like our model membrane and every biological membrane.23,24 To model the mechanical behavior of the outer membrane of the Gram-negative cell wall, the pure phospholipid membrane had to be modified by coating the membrane surface with a crystalline layer, here with streptavidin. Because the streptavidin and phospholipid layers were strongly coupled by biotin-streptavidin bonds, they formed together a compound material with mechanical properties such as bending rigidity, shear stiffness, :: :: (23) Dobereiner, H. G.; Kas, J.; Noppl, D.; Sprenger, I.; Sackmann, E. Biophys. J. 1993, 65, 1396–1403. (24) McMahon, H. T.; Gallop, J. L. Nature (London) 2005, 438, 590–596.
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and elastic area dilatation module differing from the individual layers. For pure phospholipid membranes, the bending rigidity varies in a relatively low range from 0.5 to 1 10-19 J.25,26 The bending rigidity of a membrane tethered by streptavidin is expected to be much higher than that of a pure phospholipid membrane; however, no value is known in the literature. The elastic area dilatation moduli of phospholipid membranes cover a range of 100-300 mN/m,10,25 while the elastic area dilatation of streptavidin tethered membrane is approximately 2-3 times lower (68 ( 27 mN/m).10 Therefore, instead of dynamic and reversible membrane fluctuation and budding, the rigid compound membrane alone relaxed inelastically and irreversible by surface folding and wrinkling with local high energy curvatures and, presumably, in plane shear deformations during dehydration (Figure 4, STR+GUV). In contrast to streptavidin coated GUVs, double-shell vesicles also coated with streptavidin underwent shape transformations like membrane budding without surface wrinkling of the streptavidin shell up to 700 mosm/L hyperosmotic pressures. This marked difference between the osmotic behavior of unilamellar and double-shell vesicles could be explained by the following hypothesis. In the system presented here, osmotic response is dominated by processes occurring immediately after transfer of the vesicles into hyperosmotic solution because the driving forces for deformation are highest during this initial phase. Osmotically driven flow of water through the membranes is governed by the following equation: 1 dV ¼ -Pf vw Δc A dt
ð2Þ
where A denotes membrane area, Pf is the permeability coefficient for water, vw is the molar volume of water, and Δc is the osmolarity difference acting across this membrane. For the inner membrane, V stands for the enclosed volume, and for the outer membrane V denotes the volume of the gap in between. Similar equations hold for the permeation of liquid membranes by solutes like glucose. However, permeability coefficients for water are in the range of 10-4-10-5 m/s,27-29 whereas the permeability coefficient of glucose is in the order of 10-9 m/s.30 Due to this dramatic difference in permeabilities, we can focus on water transport alone. Moreover, the osmolarity difference Δc in eq 2 changes only via volume changes. This results in characteristic response times of the compartments that are proportional to the ratio of enclosed volume to surface area.28 The decisive aspect of DSV geometry is the enormous difference in enclosed volume between the gap and inner vesicle: Vgap h ≈3 Ri Vi
ð3Þ
where Ri denotes the vesicle radius and h is the gap thickness. Based on the extremely faint fluorescence from Alexa488 enclosed (25) Rawicz, W.; Smith, B. A.; McIntosh, T. J.; Simon, S. A.; Evans, E. Biophys. J. 2000, 79, 328–339. :: (26) Dobereiner, H. G.; Gompper, G.; Haluska, C. K.; Kroll, D. M.; Petrov, P. G.; Riske, K. A. Phys. Rev. Lett. 2003, 91, 048301. (27) Huster, D.; Jin, A. J.; Arnold, K.; Gawrisch, K. Biophys. J. 1997, 73, 855– 864. (28) Olbrich, K.; Rawicz, W.; Needham, D.; Evans, E. Biophys. J. 2000, 79, 321– 327. (29) Paula, S.; Volkov, A. G.; Van Hoek, A. N.; Haines, T. H.; Deamer, D. W. Biophys. J. 1996, 70, 339–348. (30) Berglund, A. H.; Nilsson, R.; Lijenberg, C. Plant Physiol. Biochem. 1999, 37, 179–186.
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in the gap (Figure 2), a conservative upper limit for gap thickness is 100 nm. Thus, Vgap/Vi is well below 1%, probably in the range of 10-3. Therefore, the gap between the inner and outer membrane responds orders of magnitude faster to osmotic changes compared to the volume enclosed by the inner membrane. Thus, we can consider a simple steady state where the osmolarity differences over the different membranes obey Δci Pfi ¼ Δcout Pfout
ð4Þ
The permeability of the outer membrane is certainly reduced by the streptavidin crystal on top. However, its crystal structure is rather open, leaving at least 20% of each unit cell uncovered.9 Therefore, the protein crystal reduces Pfout at most to 1/5. In view of the high water content of protein crystals, an ever more modest reduction to about 1/2 appears more realistic. Taken together, this osmosis process implies a rapid, significant volume decrease of the gap region. At the same time, the inner vesicle loses volume much slower, resulting in a vigorous compression of the outer bilayer onto the inner one. The fate of the excess area in the outer bilayer depends strongly on its mechanical properties. A fluid layer (lipid bilayer alone or covered with avidin) can form buds, while the crystalline and therefore rigid outer layer of streptavidin covered DSVs must form folds like crumpled foil with areas of extremely high curvatures. However, these folds are superimposed on the spherical geometry enforced by the inner vesicle (Figures 5 and 6). The crucial assumption of this hypothesis is that such areas of high curvature act as nuclei for pore formation. At present, the role of high curvature regions for membrane fusion and fission, two processes closely related to pore formation, is intensely discussed in the context of protein mediated membrane fission or fusion.31 As in streptavidin covered DSVs, the lipid layer is tightly bonded to a rigid crystal and pore growth is inhibited, enabling efficient resealing of the lipid layer after release of excess osmotic pressure. Such resealing was observed even for fluid bilayers.32 However, the necessary temporal resolution could not be achieved with our instrumentation. This hypothesis is in accordance with all observations. It easily explains the round and relatively smooth shapes observed for streptavidin coated DSVs even at high osmotic pressures (Figure 4) and the differences between DSV, avidin coated DSV, and streptavidin covered DSV (Figure 5). Most importantly, a volume increase of the gap with a persistent encapsulation of dye in streptavidin coated DSVs, as presented in Figure 6, can only be explained by an inflow of glucose through transient pores. Unfortunately, we were not able to observe such transient pores directly. This was prevented by the very small amount of dye that can be trapped in the narrow gap between the inner and outer membranes combined with lack of time resolution of our microscope and the small size of the high curvature regions. At present, it is unclear to what extent active adaptation processes of Gram-negative bacteria can protect against osmotic stress and at what osmotic pressure mechanical processes as observed here set in. In several publications of Gervais and coworkers,3,4,33 the molecular regulatory pathway of E. coli was reported to balance turgor pressure in a hyperosmotic pressure range of 0-10 MPa, which corresponds to 0-4.4 osm/L. The mechanical pathway was first activated above 4.4 osm/L. However, Koch and Schwarz published electron microscopic images of (31) Chernomordik, L. V.; Kozlov, M. M. Nat. Struct. Mol. Biol. 2008, 15, 675– 683. (32) Riske, K. A.; Dimova, R. Biophys. J. 2005, 88, 1143–1155. (33) Beney, L.; Mille, Y.; Gervais, P. Biophys. J. 1997, 72, 1258–1263.
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E. coli exposed to hyperosmotic pressure of 500 mosm/L.7 These pictures clearly showed the typical signals of mechanical response such as cytoplasmic membrane budding and surface wrinkling of the outer membrane complex even under this comparatively gentle dehydration. Because we did not study living cells, this open question cannot be answered here. Nevertheless, based on the successful mimicking of the complete mechanical process using double-shell giant vesicles, we could demonstrate that the encapsulated membrane structure very effectively protects the
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membrane under high osmotic pressures even in the absence of active cellular processes. These findings support the hypotheses that the double-membrane architecture of Gram-negative cell walls plays a vital role in the osmotic regulatory system and that it is highly efficient during extreme dehydration. Judged from our biomimetic model system, this purely mechanical protection could compensate an additional pressure range of 2-5 MPa without any support of an active molecular regulatory pathway.
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