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Polymer Vesicles as Microreactors for Bioinspired Calcium Carbonate Precipitation Andreas Picker,† Hermann Nuss, Patrick Guenoun, and Corinne Chevallard* IRAMIS, LIONS, UMR SIS2M 3299 CEA-CNRS, CEA-Saclay, F-91191 Gif-sur-Yvette Cedex, France
bS Supporting Information ABSTRACT: Giant polymer vesicles made by electroformation have been shown to encapsulate salts up to concentrations of about 10 mM. The impermeability of these “polymersomes” to calcium ions is demonstrated by the use of fluorescent probes dedicated to calcium analysis. Permeability to calcium ions can be triggered by the addition of calcimycin, an ionophore molecule that is able to transport cations selectively through the membrane. As a result, we show that the mineralization of calcium carbonate can be induced within the polymersomes, which were previously loaded with carbonate ions. This is a further step toward the use of polymersomes as microreactors and the study of mineralization schemes, including biomimetic ones, in confined environments.
’ INTRODUCTION 1. Introduction. Biominerals are biogenic materials made by inorganic precipitation in contact with some organic matter. They form under the cellular control of a living organism. In several cases, it is believed that the inorganic phase, in unprecipitated or amorphous form, is conveyed to the site of mineralization by lipid vesicles, where precipitation or crystallization takes place.1,2 In nacre mineralization, calcium carbonate might precipitate first as an amorphous phase in intracellular vesicles before crystallizing near or in contact with an insoluble extracellular organic matrix. Nacre mineralization may also occur from mineral precipitation within the closed compartments delineated by this organic scaffold. Therefore, synthetically produced vesicles may be viewed as model confined environments of tunable size and chemistry that are appropriate for studying the influence of space confinement on biomineralization processes. These remarks have generated a number of studies consisting of precipitating various minerals inside organic vesicles, which are mostly lipidic.3-8 A key point is then to control the permeation of the reactants through the vesicle membrane in order to define the place and time of precipitation. This control can be based on the difference in permeation properties between some intravesicular cations, which are assumed to not leak out, and an extravesicular anion, such as hydroxyde, that is able to penetrate the vesicle and trigger the precipitation of various metallic minerals.3-6 Other experiments were carried out by Eanes and co-workers on calcium phosphate precipitation that achieved kinetic control by inducing a permeability of the vesicles to external calcium ions, thanks to a cationophore such as lasalocid acid.7 These experiments underlined the prevalence of mineral nucleation on the internal walls of the vesicles. Temperature was used as r 2011 American Chemical Society
another effective trigger for calcium permeation when vesicles were made of lipids close to their chain-melting transition.8 A quantitative evaluation of the permeability to calcium ions was performed for liposomes through the use of a synthetic polymer (poly(2-ethylacrylic acid) that induced channel formation within the membrane.9 In this letter, we make use of polymer vesicles, so-called “polymersomes”, to control the encapsulation and precipitation of calcium carbonate. Polymersomes, made by the self-assembly of block copolymers, are advantageous over lipid vesicles because they are usually mechanically tougher, more impermeable to both cations and anions, and more versatile in terms of the chemical nature of their hydrophilic and hydrophobic moieties.10,11 The two latter aspects are particularly important because they provide ways to induce polymersome permeability to ions by applying an external trigger such as the addition of selective or unselective ion channel molecules to the membrane.12 Also, a large variety of copolymers have now been used to make polymersomes, which provides the possibility to vary the bare vesicle permeability or the nature of the precipitates formed by heterogeneous nucleation at the inner wall of the polymersome. In particular, the hydrophilic moiety of the polymer can be selected to mimic at best the organic matrix of Nacre that is covered with adsorbed acidic proteins.13 Recent studies have succeeded in inducing permeability into stimuli-responsive polymersomes, thus releasing encapsulated fluorescent molecules or enzymes14 or enabling ions to penetrate polymersomes by the use of various ionophores.12 However, quantitative measurements of polymersome permeability have Received: September 21, 2010 Revised: January 19, 2011 Published: March 04, 2011 3213
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been limited to water permeation induced by an osmotic shock10,15-17 or passive permeation in the specific case of 5,50 -dithiobis-(2-nitrobenzoic acid).18 None of these permeability studies provided in situ followup for the polymersome population and permeability heterogeneities. In our study, giant polymersomes reminiscent of biomineralized organic compartments were produced by electroformation, and the efficiency of this technique with respect to calcium and carbonate ion encapsulation was systematically investigated. The bare and induced permeabilities of the electroformed polymersomes to calcium ions were investigated by visual inspection and measured by using ion-sensitive fluorescent dyes. The polymersomes were subsequently used as microreactors for inner calcium carbonate growth, and the mineral growth was observed in situ.
’ MATERIALS AND METHODS Materials. Chemicals. Calcium chloride dihydrate (CaCl2 3 2H2O, >99.5% purity) was purchased from Fluka, and sodium carbonate (Na2CO3 > 99% purity) was purchased from Labosi. Both were used as received. Chloroform (Riedel-de Ha€en), ethanol (Merck), and acetonitrile (SDS) solvents were all analytical grade. The water used was ultrapure (18.2 MΩ 3 cm, pH 5.6), produced by a Millipore filtration system. Polymer. The polymer used to form polymer vesicles is a comblike copolymer, named PEG-12 dimethicone, that has 2 PEG arms having 12 monomers on average that are randomly distributed on a PDMS backbone. This product, of formula (CH3)3SiO(Si(CH3)2O)22(Si(CH3)(M)O)2Si(CH3)3 with M = (CH2)3(OCH2CH2)12OH, is synthesized and commercialized by Dow Corning under the reference Dow 5329.19 Its molecular weight is estimated to about 3000 g 3 mol-1 from bulk melt viscosity measurements with a high polydispersity.20,21 In view of electroformation, Dow polymer was dissolved in chloroform with 5% v/v acetonitrile to a final concentration of 10 mg 3 mL-1. Calcium-Sensitive Dyes. Calcium-sensitive fluorophores Fluo-3 (F1240) and Fura-2 (F-1200) were purchased from Invitrogen. Fluo-3 is a single-wavelength dye with a maximum excitation wavelength of 506 nm and an emission wavelength of 526 nm. Its fluorescence is almost zero in the absence of calcium but increases gradually upon calcium binding (Supporting Information). In contrast, Fura-2 is a ratiometric dye whose excitation and emission spectra change in response to calcium binding. More precisely, its excitation peak moves from λ = 363 nm in the absence of calcium to λ = 335 nm in response to calcium binding. As shown in the Supporting Information, it is effectively sensitive to calcium concentration changes in the range of 1-10 μM. Carrier Ionophores. Carrier ionophore calcimycin (A23187) was purchased from Invitrogen and dissolved in DMSO to prepare a 10 mM stock solution. It selectively binds cations, with a much higher selectivity for divalent cations (Mn2þ . Ca2þ ≈ Mg2þ . Sr2þ > Ba2þ) than for monovalent ions (Liþ > Naþ > Kþ).22 It can be used to equilibrate the cation concentrations quickly on either side of a lipid membrane because it forms a neutral complex with the cations and can then diffuse through the membrane. It has mainly been used to investigate calcium-regulated cellular mechanisms.23-25 Polymer Vesicle Electroformation. To form giant vesicles, we adapted the electroformation technique devised by Angelova et al.26 This technique initially developed for lipids has recently been used to form polymer vesicles from diblock27,28 or triblock12 copolymers. More precisely, we used the method described in Mathivet et al.29 Conductive glass slides, exhibiting a transparent indium-tin oxide (ITO) layer, are spin-coated with a Dow polymer film at room temperature. The optimal thickness of the film with respect to the electroformation efficiency is provided by a rotation speed of 600 rpm, in total agreement with the conclusion drawn by Estes et al. in the case of lipids.30 A 2 V, 10 Hz sine wave is applied between the
Figure 1. Vesicles developed on the ITO-coated slides upon electroformation at different calcium salt concentrations. two ITO slides coated with the polymer film, which enclose the electroformation chamber. Then, the latter is filled with water or with the desired salt solution. Once vesicle growth is achieved, the release of the vesicles in the bulk is promoted by applying a square-wave voltage of 2 V at 5 Hz for 10 min, which is then lowered to 2 Hz for 10 min. In addition, a gentle shear flow is applied to pour the vesicles out of the chamber, and it helps to detach quite a few of them. Dialysis Protocols. Dialysis of the vesicle solutions was achieved in SpectraPor dialysis bags with a molecular weight cutoff of 8 kDa to remove salt ions and 100 or 300 kDa to remove fluorophore molecules. Dialysis to remove salt ions was performed against glucose solutions under isotonic conditions, with some NaN3 added (0.01 wt %) to prevent bacteria formation. Characterization. Fluorescence images were captured using a confocal fluorescence microscope (Olympus IX81 inverted microscope with an Olympus Fluoview-1000 confocal head), whereas phase-contrast microscopy was performed on an Olympus upright BX61WI microscope. Finally, fluorometry measurements were made on a Varian Cary Eclipse fluorometer equipped with a Xe lamp.
’ RESULTS AND DISCUSSION Calcium and Carbonate Encapsulation. The electroformation technique consists of forming vesicles by applying a lowfrequency sinusoidal electric field to a lipid or polymer film. In the case of neutral species, it is generally believed that the electroosmotic flow induced by this ac electric field results in the mechanical shearing of the swollen multilayer film, which induces vesicle formation through bilayer destabilization.31 For neutral lipids, such a process has been reported to be successful for ionic strengths lower than 10 mM.31,32 At higher ionic strengths, it is likely that the screening of the electric field reduces the amplitude of the electro-osmosis effect and therefore impedes vesicle formation. Still, a few publications have reported the successful electroformation of polymer vesicles in buffer solutions.12,28 We therefore decided to probe the range of efficiency of the electroformation technique with respect to calcium and carbonate salt encapsulation. Electroformation with calcium chloride (CaCl2) solutions was successfully performed for concentrations ranging from 0.1 to 8 mM (Figure 1), whereas vesicles were never observed for concentrations higher than 8 mM. Moreover, the 3214
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Figure 2. Result of vesicle electroformation in a 2 mM Na2CO3 solution (duration: 160 min).
higher the salt concentration, the longer the time needed to obtain polymer vesicles under similar spin-coating conditions. Whereas vesicles appear almost instantaneously in water, it takes 60 to 120 min to form vesicles at 4 mM CaCl2. With increasing salt concentration, the efficiency of the vesicle-formation process becomes more and more sensitive to the thickness homogeneity of the polymer film such that any defect in the layer may result in poor vesicle formation. Consequently, thickness heterogeneities result in ITO slide regions exhibiting no vesicles. Moreover, the largest diameter that can be reached in 2 mM calcium solution is about 50 μm. Electroformation in the presence of sodium carbonate (Na2CO3) could be achieved for concentrations of up to 15 mM (cf. Figure 2). No significant change in the electroformation time or typical vesicle size was noticed in comparison to the electroformation in a calcium chloride solution. A systematic study of the vesicle stability with time showed that vesicles, when electroformed in the presence of salt, are stable for at least 1 week (Supporting Information). To probe the permeability properties of the Dow polymer vesicles with respect to calcium ions, vesicles were formed in a 5 μM solution of calcium-sensitive dye Fluo-3. Whereas Fluo-3 scarcely fluoresces in the absence of calcium, it exhibits a typical 100-fold increase upon calcium binding.33 Moreover, its low dissociation constant Kd makes it sensitive to calcium concentrations of as low as 20 nM.34 A CaCl2 solution was subsequently added to the outer medium up to a final concentration of 10 μM.35 Figure 3a shows vesicles that remained nonfluorescent upon calcium addition, which demonstrates their impermeability to calcium ions. However, one must underline that not all vesicles exhibit such behavior and that some do show an increase in fluorescence upon calcium addition. It is likely that this originates from the inhomogeneity of the vesicle population, which may contain vesicles with pores according to the way that they detached from the polymer layer during electroformation. Nonetheless, for impermeable vesicles, observations made more than 2 weeks after salt addition could provide evidence that a majority of vesicles were still impermeable to calcium ions. The encapsulation efficiency of the electroformation technique could be determined using ratiometric dye Fura-2. Vesicles containing calcium salt solution in the micromolar range and 10 μM Fura-2 were dialyzed against water.36 Contrary to single excitation dyes such as Fluo-3, ratiometric dyes such as Fura-2 allow one to measure the intravesicular calcium concentration in a way that is not sensitive to the fluorophore concentration and therefore to bleaching effects. The Ca2þ concentration is deduced from the measurement of the ratio R between the fluorescence intensity values that correspond to two excitation wavelengths, λ1 = 340 nm and λ2 = 380 nm. Using a calibration curve relating R to [Ca2þ] (see Figure 3c and the
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Supporting Information for its construction), one can then deduce the unknown concentration of calcium from the measured R value, provided that the calcium concentration lies within the sensitivity range of the dye (1 to 10 μM). This procedure was followed for vesicles electroformed in solutions of 6 and 10 μM Ca2þ (Figure 3b). The measured R value, 3.2 in the former case and 4.9 in the latter, corresponds to concentrations deduced from the calibration curve of about 5.7 ( 0.6 and 10.5 ( 1.0 μM, respectively. These results confirm the efficiency of the electroformation technique in trapping calcium salt within vesicles. They also prove the impermeability of the polymer vesicles to calcium ions after dialysis against water for 3 days. Ionophore-Induced Permeability of the Polymersomes to Calcium Ions. Calcimycin is a well-known ion carrier that has been used in many studies to provide calcium permeability to lipid vesicles37,38 or cells.39 However, to our knowledge calcimycin had never been used with polymersomes; therefore, it was questionable whether it could diffuse through the polymersome membrane and efficiently carry calcium ions through the membrane. Depending on its relative concentration with respect to lipid concentration, calcimycin has been shown to behave either as an ion carrier, similar to ionophores Lasalocid A and N,Ndicyclohexyl-N0 ,N0 -dioctadecyl-3-oxapentane-1,5-diamide used by Sauer et al.12 to provide calcium permeability to PDMS-based triblock copolymer vesicles, or as an ion channel.40 The latter behavior leads to much faster ion diffusion through the membrane than the former. We therefore made polymersomes containing 5 μM Fluo-3 dye and dialyzed them against water. We then added Ca2þ to a final concentration of 20 μM. As shown in Figure 4a, no change in the fluorescence signal of Fluo-3 occurs upon calcium addition, confirming that calcium ions cannot enter polymersomes. The addition of the ionophore to a final concentration of 10 μM in the extravesicular medium leads to a huge increase in the fluorescence intensity, which expresses the influx of calcium ions into the vesicles. However, as the ionophore itself fluoresces in the absence of calcium, this experiment was repeated by adding first the ionophore and then calcium ions. In this case, the fluorescence signal increases after calcimycin addition (Figure 4b) and saturates after a while (Figure 4b,c). Adding calcium salt then induces a second increase in the fluorescence signal associated with the calcium influx in the vesicle. For the chosen concentration of ionophore, this influx occurs on a relatively long timescale and the fluorescence signal saturates after about 30 min (Figure 4c). Assuming that the final (at time t = 30 min) intravesicular concentration is equal to the outer concentration (20 μM) and that the average vesicle diameter is 10 μm, one can estimate a typical membrane permeability P using the relation J = PSΔC, where J is the molar flux of calcium ions, S is the surface area of the vesicle, and ΔC is the calcium concentration difference between the intra- and extravesicular media. This gives P ≈ 1 nm 3 s-1, which is the same order of magnitude as the permeability of polymer membranes to polar molecules given in ref 18. Increasing the ionophore concentration up to 40 μM did not speed up the calcium permeation significantly (data not shown). Calcium Carbonate Precipitation. The previous experiment was repeated on carbonate-containing vesicles in order to induce intravesicular CaCO3 precipitation. Carbonate-containing vesicles were produced using electroformation in a 4 mM Na2CO3 solution. Dialysis was performed against a glucose solution to remove the extravesicular carbonate salt without inducing strong osmotic effects. We then added 10 mM calcium and 10 μM calcimycin to the outer medium. Within a few minutes after this addition, 3215
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Figure 3. (a) Fluorescence image of Fluo-3-containing vesicles after Ca2þ addition to a final concentration of 10 μM. (b) Excitation spectra of Fura-2 within electroformed vesicles at an emission wavelength of λ = 510 nm. The left spectrum corresponds to vesicles formed in a 6 μM CaCl2 solution, and the right spectrum corresponds to vesicles formed in a 10 μM solution. (c) Calibration curve of Fura-2 relating R = F340/F380 to [Ca2þ].
precipitation in polymersomes becomes visible in the form of many black spots mostly located at the inner walls of the vesicles, as evidenced by phase-contrast microscopy (Figure 5). The number of vesicles exhibiting precipitation increased over time, and the quantity of precipitates appeared to saturate after several tens of minutes. A more precise quantification of the mineral growth kinetics was not achieved because of the variability in the precipitation kinetics from one vesicle to the other. In a comparative experiment, only calcium and no calcimycin was added to a carbonate-containing vesicle solution; the vesicles
showed shape modifications due to the induced osmotic shock, but without precipitation inside the vesicles (pictures not shown). These black spots observed under the polarizing microscope did not show any birefringence, indicating the amorphous character of the calcium carbonate mineral. Preliminary infrared spectroscopy experiments could provide further evidence that the precipitation is indeed due to CaCO3 formation in spite of the very low quantity of material. However, assuming that all of the encapsulated carbonate ions are precipitated, the expected 3216
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Figure 4. (a) Fluorescence emission spectra of Fluo-3-containing vesicles after the successive addition of 20 μM calcium ions and 10 μM calcimycin. (b) Fluorescence emission spectra of Fluo-3-containing vesicles after the sequential addition of calcimycin and calcium salt. The curve that shows calcimycin fluorescence was recorded after the fluorescence increase had reached a maximum, that is, after full diffusion of the ionophore in the solution. (c) Increase in the maximum fluorescence intensity with respect to the time elapsed since calcium addition (curve deduced from the spectra in b). The solid black curve is just a guide for the eyes.
Figure 5. Intravesicular CaCO3 precipitation induced by calcimycin addition.
volume of mineral correspond to ∼0.15 μm3 or alternatively to a single mineral particle of diameter in the range of 530-650 nm.41 Such a calculation is in reasonable agreement with disklike precipitates formed at the membrane, possibly as a result of heterogeneous precipitation. Moreover, this experiment points out the low permeability of the polymersomes to carbonate ions because vesicles dialyzed against water for up to 6 days still exhibited precipitation.
’ CONCLUSIONS To our knowledge, this letter is the first report of calcium carbonate precipitation induced within polymer vesicles. This intravesicular precipitation was triggered by the calcimycin ionophore, which in turn induced a calcium influx into carbonatecontaining giant polymersomes. The kinetics of this calcium influx needs to be investigated further and can probably be finely tuned by changing the concentration or the nature of the ionophore. Furthermore, the current development of microfluidic tools ensuring the production of giant unilamellar vesicles with a high size monodispersity15 should provide better control of the vesicle properties, and therefore a higher reproducibility of the encapsulation and precipitation results. In any case, this work opens the way to the use of polymersomes as efficient microreactors. Their versatile size and chemistry will be used to gain a better understanding of the mineral nucleation triggered within organic compartments, which is a major feature of biomineralization. ’ ASSOCIATED CONTENT
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Supporting Information. Time and shape stability of vesicles. Fluorescence spectra of calcium-sensitive dye Fluo-3.
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’ AUTHOR INFORMATION Corresponding Author
*Phone: þ33 (0)1 69 08 52 23. Fax: þ33 (0)1 69 08 66 40. E-mail:
[email protected]. Present Addresses †
Department of Chemistry, POB 714, Universit€at Konstanz, D-78457 Konstanz, Germany.
’ ACKNOWLEDGMENT This work was supported by the European Master “Complex Condensed Materials and Soft Matter” (EMASCO-COSOM) and by the French research program ANR-07-NANO-061-01 “Monopoly”. We thank A. Carlsen, J.-F. Le Meins, C. Schatz, and S. Lecommandoux (LCPO/ENSCPB, Bordeaux) for fruitful scientific discussions and for the gift of Dow 5329 polymer. We also acknowledge M. Le Maire (CEA/DSV) for his advice concerning the use of ionophore A23187. ’ REFERENCES (1) Young, J. R.; Henriksen, K. Rev. Mineralo. Geochem. 2003, 54, 189–215. (2) Eanes, E. D.; Heywood, B. R. Liposome Technology in Biomineralization Research. New Biotechnology in Oral Research; Meyers, H. M., Ed.; Karger: Basel, Switzerland, 1989; pp 54-75. (3) Mann, S.; Williams, R. J. P. J. Chem. Soc., Dalton Trans. 1983, 2, 311–316. (4) Mann, S.; Hannington, J. P. J. Colloid Interface Sci. 1987, 122, 326–335. (5) Bhandarkar, S.; Bose, A. J. Colloid Interface Sci. 1990, 135, 531–538. (6) Yaacob, I. I.; Bhandarkar, S.; Bose, A. J. Mater. Res. 1993, 8, 573–577. (7) Eanes, E. D.; Hailer, A. W.; Costa, J. L. Calcif. Tissue Int. 1984, 36, 421–430. (8) Messersmith, P. B.; Starke, S. Chem. Mater. 1998, 10, 117–124. (9) Thomas, J. L.; Tirrell, D. A. J. Controlled Release 2000, 67, 203–209. (10) Discher, B. M.; Won, Y. Y.; Ege, D. S.; Lee, J. C. M.; Bates, F. S.; Discher, D. S.; Hammer, D. A. Science 1999, 294, 1143–1146. (11) LoPresti, C.; Lomas, H.; Massignani, M.; Smart, T.; Battaglia, G. J. Mater. Chem. 2009, 19, 3576–3590. (12) Sauer, M.; Haefele, T.; Graff, A.; Nardin, C.; Meier, W. Chem. Commun. 2001, 23, 2452–2453. (13) Cartwright, J. H. E.; Checa, A. G. J. R. Soc., Interface 2007, 4, 491–504. (14) Kim, K. T.; Cornelissen, J. J. L. M.; Nolte, R. J. M.; van Hest, J. C. M. Adv. Mater. 2009, 21, 2787–2791. (15) Lorenceau, E.; Utada, A. S.; Link, D. R; Cristobal, G.; Joanicot, M.; Weitz, D. A. Langmuir 2005, 21, 9183–9186. (16) Kumar, M.; Grzelakowski, M.; Zilles, J.; Clark, M.; Meier, W. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 20719–20724. (17) Sanson, C.; Schatz, C.; Le Meins, J. F.; Br^ulet, A.; Soum, A.; Lecommandoux, S. Langmuir 2009, 26, 2751–2760. (18) Battaglia, G.; Ryan, A. J.; Tomas, S. Langmuir 2006, 22, 4910–4913. (19) The given formula represents only the average structure. Moreover, according to ref 14, Dow 5329 exhibits 85% purity. The precise polydispersity is unknown. (20) Lin, Z.; Hill, R. M.; Davis, H. T.; Scriven, L. E.; Talmon, Y. Langmuir 1994, 10, 1008–1011.
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(21) Nam, J.; Santore, M. M. Langmuir 2007, 23, 7216–7224. (22) Pfeiffer, D. R.; Reed, P. W.; Lardy, H. A. Biochemistry 1974, 13, 4007–4014. (23) Murtha, A. P.; Feng, L. P.; Grotegut, C.; Schomberg, D. Reprod. Sci. 2009, 16, 295A–296A. (24) Doyle, A. D.; Lee, J. J. Cell Sci. 2005, 118, 369–379. (25) Mao, Y.; Xiang, H.; Wang, J.; Li, D. Investig. Ophthalmol. Visual Sci. 2002, 43, U842. (26) Angelova, M. I.; Dimitrov, D. S. Faraday Discuss. Chem. Soc. 1986, 81, 303–311. (27) Dimova, R.; Seifert, U.; Pouligny, B.; F€orster, S.; D€ obereiner, H.-G. Eur. Phys. J. E 2002, 7, 241–250. (28) Lomas, H.; Massignani, M.; Abdullah, K. A.; Canton, I.; LoPresti, C.; MacNeil, S.; Du, J.; Blanazs, A.; Madsen, J.; Armes, S. P.; Lewis, A. L.; Battaglia, G. Faraday Discuss. 2008, 139, 143–159. (29) Mathivet, L.; Cribier, S.; Devaux, P. F. Biophys. J. 1996, 70, 1112–1121. (30) Estes, D. J.; Mayer, M. Colloids Surf., B 2005, 42, 115–123. (31) Angelova, M. I.; Dimitrov, D. S. Prog. Colloid Polym. Sci. 1988, 76, 59–67. (32) Bucher, P.; Fischer, A.; Luisi, P. L.; Oberholzer, T.; Walde, P. Langmuir 1998, 14, 2712–2721. (33) Harkins, A. B.; Kurebayashi, N.; Baylor, S. M. Biophys. J. 1993, 65, 865–881. (34) http://www.invitrogen.com/site/us/en/home/References/ Molecular-Probes-The-Handbook/Indicators-for-Ca2-Mg2-Zn2-andOther-Metal-Ions/Fluorescent-Ca2-Indicators-Excited-with-VisibleLight.html (35) Such a low concentration does not induce any significant osmotic pressure (Π = 0.000734 atm). Moreover, the impermeability of the vesicles to Fluo-3 was checked independently. (36) The impermeability of the vesicles to Fura-2 has been checked independently. (37) Erdahl, W. L.; Chapman, C. J.; Taylor, R. W.; Pfeiffer, D. R. Biophys. J. 1994, 66, 1678–1693. (38) Wang, E.; Taylor, R. W.; Pfeiffer, D. R. Biophys. J. 1998, 75, 1244–1254. (39) Drummond, I. A. S.; Lee, A. S.; Resendez, E.; Steinhardt, R. A. J. Biol. Chem. 1987, 262, 12801–12805. (40) Jyothi, G.; Surolia, A.; Easwaran, K. R. K. J. Biosci. 1994, 19, 277–282. (41) The lower diameter value corresponds to the calcite mass density (2.71 g 3 cm-3) whereas the larger corresponds to the amorphous phase of density 1.49 g 3 cm-3 reported in Bolze, J.; Peng, B.; Dingenouts, N.; Panine, P.; Narayanan, T.; Ballauff, M. Langmuir 2002, 18, 8364–8369.
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