Kinetic and Structural Characterization of Adsorption-induced Unfolding of Bovine α-Lactalbumin
2002; Elsevier BV; Volume: 277; Issue: 13 Linguagem: Inglês
10.1074/jbc.m106005200
ISSN1083-351X
AutoresMaarten F. M. Engel, Carlo P. M. van Mierlo, Antonie J. W. G. Visser,
Tópico(s)Protein purification and stability
ResumoConformational changes of bovine α-lactalbumin induced by adsorption on a hydrophobic interface are studied by fluorescence and circular dichroism spectroscopy. Adsorption of bovine α-lactalbumin on hydrophobic polystyrene nanospheres induces a non-native state of the protein, which is characterized by preserved secondary structure, lost tertiary structure, and release of calcium. This partially denatured state therefore resembles a molten globule state, which is an intermediate in the folding of bovine α-lactalbumin. Stopped-flow fluorescence spectroscopy reveals two kinetic phases during adsorption with rate constants k1 ∼ 50 s−1 and k2 ∼ 8 s−1. The rate of partial unfolding is remarkably fast and even faster than unfolding induced by the addition of 5.4 mguanidinium hydrochloride to native α-lactalbumin. The large unfolding rates exclude the possibility that unfolding of bovine α-lactalbumin to the intermediate state occurs before adsorption takes place. Stopped-flow fluorescence anisotropy experiments show that adsorption of bovine α-lactalbumin on polystyrene nanospheres occurs within the dead time (15 ms) of the experiment. This shows that the kinetic processes as determined by stopped-flow fluorescence spectroscopy are not affected by diffusion or association processes but are solely caused by unfolding of bovine α-lactalbumin induced by adsorption on the polystyrene surface. A scheme is presented that incorporates the results obtained and describes the adsorption of bovine α-lactalbumin. Conformational changes of bovine α-lactalbumin induced by adsorption on a hydrophobic interface are studied by fluorescence and circular dichroism spectroscopy. Adsorption of bovine α-lactalbumin on hydrophobic polystyrene nanospheres induces a non-native state of the protein, which is characterized by preserved secondary structure, lost tertiary structure, and release of calcium. This partially denatured state therefore resembles a molten globule state, which is an intermediate in the folding of bovine α-lactalbumin. Stopped-flow fluorescence spectroscopy reveals two kinetic phases during adsorption with rate constants k1 ∼ 50 s−1 and k2 ∼ 8 s−1. The rate of partial unfolding is remarkably fast and even faster than unfolding induced by the addition of 5.4 mguanidinium hydrochloride to native α-lactalbumin. The large unfolding rates exclude the possibility that unfolding of bovine α-lactalbumin to the intermediate state occurs before adsorption takes place. Stopped-flow fluorescence anisotropy experiments show that adsorption of bovine α-lactalbumin on polystyrene nanospheres occurs within the dead time (15 ms) of the experiment. This shows that the kinetic processes as determined by stopped-flow fluorescence spectroscopy are not affected by diffusion or association processes but are solely caused by unfolding of bovine α-lactalbumin induced by adsorption on the polystyrene surface. A scheme is presented that incorporates the results obtained and describes the adsorption of bovine α-lactalbumin. Adsorption of proteins on solid/liquid interfaces is a generally occurring phenomenon both in nature and in man-made systems. It is well recognized that proteins undergo conformational changes upon adsorption on solid/liquid interfaces (1.Haynes C.A. Norde W. J. Colloid Interface Sci. 1995; 169: 313-328Crossref Scopus (352) Google Scholar, 2.Andrade J.D. Hlady V. Adv. Polym. Sci. 1986; 79: 1-63Crossref Google Scholar, 3.Baszkin A. Norde W. Physical Chemistry of Biological Interfaces. Marcel Dekker Inc., New York2000Google Scholar). Because unfolding of proteins can have a large effect on their function or properties, it is important to increase the knowledge about protein adsorption and about the resulting conformational changes. It is necessary to study not only the conformation of a protein in the adsorbed state but also the kinetics of the adsorption-induced conformational changes. This knowledge will help in controlling wanted or unwanted conformational changes of adsorbed proteins. In this work we focus on the kinetics of adsorption-induced conformational changes of bovine α-lactalbumin (BLA) 1The abbreviations used are: BLAbovine α-lactalbuminGdnHClguanidinium hydrochlorideMGmolten globuleCDcircular dichroismICP-OESinductively coupled plasma-optical emission spectrometry 1The abbreviations used are: BLAbovine α-lactalbuminGdnHClguanidinium hydrochlorideMGmolten globuleCDcircular dichroismICP-OESinductively coupled plasma-optical emission spectrometry upon interaction with colloidal polystyrene nanospheres. Detailed information on adsorption-induced conformational changes and especially on the kinetics of adsorption-induced conformational changes is sparse. An important reason for this is the experimental difficulty of the presence of an adsorbent, which is usually a solid phase. A few routes have been designed to remove this difficulty. A macroscopic solid/water interface can be used in combination with a reflective technique like ellipsometry or total internal reflection fluorescence (4.Wahlgren M.C. Arnebrant T. Paulsson M.A. J. Colloid Interface Sci. 1993; 158: 46-53Crossref Scopus (38) Google Scholar, 5.van Wagenen R. Rockhold S. Andrade J. Cooper S.L. Peppas N.A. Biomaterials: Interfacial Phenomena and Applications. 199. American Chemical Society, Washington D.C.1982: 351-370Google Scholar). In other cases a microscopic system has been used that consists of small particles with diameters ranging from nanometers to micrometers in size (6.Maste M.C.L. Pap E.H.W. van Hoek A. Norde W. Visser A.J.W.G. J. Colloid Interface Sci. 1996; 180: 632-633Crossref Scopus (40) Google Scholar, 7.Norde W. Favier J.P. Colloids Surf. 1992; 64: 87-93Crossref Scopus (362) Google Scholar). The latter system has the advantage of providing a large interface, however, light scattering and light absorbance often interfere with several spectroscopic techniques. Here, we use a suspension of polystyrene nanospheres with a diameter of ∼50 nm. The large surface area per gram of polystyrene and therefore the high binding capacity allows a reduction of the nanosphere concentration. As a result, light scattering and absorption are reduced and thus the use of spectroscopic methods like stopped-flow fluorescence is feasible. bovine α-lactalbumin guanidinium hydrochloride molten globule circular dichroism inductively coupled plasma-optical emission spectrometry bovine α-lactalbumin guanidinium hydrochloride molten globule circular dichroism inductively coupled plasma-optical emission spectrometry Bovine α-lactalbumin is a small protein (14 kDa), containing four tryptophan residues and four disulfide bonds. It can bind calcium, which plays an important role in the stability and folding behavior of the protein (8.Troullier A. Reinstadler D. Dupont Y. Naumann D. Forge V. Nat. Struct. Biol. 2000; 7: 78-86Crossref PubMed Scopus (109) Google Scholar). BLA is chosen in this study for several reasons. First, the structure, stability, and folding behavior of BLA have been thoroughly studied (9.Chrysina E.D. Brew K. Acharya K.R. J. Biol. Chem. 2000; 275: 37021-37029Abstract Full Text Full Text PDF PubMed Scopus (212) Google Scholar, 10.Greene L.H. Grobler J.A. Malinovskii V.A. Tian J. Acharya K.R. Brew K. Protein Eng. 1999; 12: 581-587Crossref PubMed Scopus (38) Google Scholar, 11.Kuwajima K. Hiraoka Y. Ikeguchi M. Sugai S. Biochemistry. 1985; 24: 874-881Crossref PubMed Scopus (288) Google Scholar). Second, BLA is a protein that is applied at solid/water interfaces, for example in food applications. Finally, the folding pathway of BLA includes a stable intermediate, the so-called molten globule state, which has been studied in detail previously (12.Vanderheeren G. Hanssens I. J. Biol. Chem. 1994; 269: 7090-7094Abstract Full Text PDF PubMed Google Scholar, 13.Kuwajima K. FASEB J. 1996; 10: 102-109Crossref PubMed Scopus (430) Google Scholar, 14.Kim S. Bracken C. Baum J. J. Mol. Biol. 1999; 294: 551-560Crossref PubMed Scopus (38) Google Scholar). Isothermal titration calorimetry and intrinsic fluorescence spectroscopy have indicated denaturation of BLA upon adsorption on hydrophobic interfaces (15.Haynes C.A. Sliwinsky E. Norde W. J. Colloid Interface Sci. 1994; 164: 394-409Crossref Scopus (156) Google Scholar, 16.Oroszlan P. Blanco R. Lu X.M. Yarmush D. Karger B.L. J. Chromatogr. 1990; 500: 481-502Crossref PubMed Scopus (73) Google Scholar). A study of Banuelos and Muga (17.Banuelos S. Muga A. J. Biol. Chem. 1995; 270: 29910-29915Abstract Full Text Full Text PDF PubMed Scopus (104) Google Scholar) shows that BLA binds to lipid bilayers in a molten globule-like conformation. Recently some studies (18.Karlsson M. Martensson L.-G. Jonsson B.-H. Carlsson U. Langmuir. 2000; 16: 8470-8479Crossref Scopus (55) Google Scholar, 19.Surrey T. Jähnig F. J. Biol. Chem. 1995; 270: 28199-28203Abstract Full Text Full Text PDF PubMed Scopus (103) Google Scholar, 20.Sanghera N. Pinheiro T.J. Protein Sci. 2000; 9: 1194-1202Crossref PubMed Scopus (72) Google Scholar) emerged that describe the kinetics of conformational changes induced by adsorption on an interface or interaction with lipid vesicles. Unfortunately, still little is known about the altered conformation in the adsorbed state and about the kinetics of the conformational changes that take place upon adsorption. Further knowledge on these matters is of particular relevance for the understanding of protein-membrane interactions and in the field of synthetic biomaterials used for medical applications. Regarding protein-membrane interactions, it is hypothesized that, when proteins are translocated across the membrane, they must be in a non-native or molten globule (MG) state (21.Ptitsyn O.B. Adv. Protein Chem. 1995; 47: 83-229Crossref PubMed Google Scholar). Previous work has indicated that, upon interaction with model membranes, BLA indeed can adopt an MG state (17.Banuelos S. Muga A. J. Biol. Chem. 1995; 270: 29910-29915Abstract Full Text Full Text PDF PubMed Scopus (104) Google Scholar). However, the interaction of proteins with model membranes can occur via two mechanisms: (a) adsorption on the membrane interface and (b) combined adsorption and insertion/penetration into the membrane bilayer (22.Hanssens I. Houthuys C. Herreman W. van Cauwelaert F.H. Biochim. Biophys. Acta. 1980; 602: 539-557Crossref PubMed Scopus (46) Google Scholar). In the study presented here, BLA is adsorbed on a solid interface, so insertion or penetration cannot occur. The results show that adsorption itself suffices to cause a conformational change of BLA to an adsorbed state with molten globule properties. Although the polystyrene nanospheres used are not a biological system, they are, at the moment, the only solid/liquid interface system that allows the study of the effect of adsorption (excluding penetration) on protein conformation with kinetic spectroscopic methods. We believe that the results obtained widen the biological view of protein-membrane interactions. Furthermore, our study is relevant in the field of synthetic biomaterials. Synthetic polymer biomaterials are used in medical applications, for example as implants in the human body. Contact between these biomaterials and tissue or blood gives rise to all kinds of complex and mostly unwanted phenomena. Upon implantation of foreign material in the human body, proteins adsorb on the biomaterial. Knowledge of this process is relevant in the search for new and better biomaterials that can reduce adverse reactions of the human body. In this report, the adsorption-induced unfolding of bovine α-lactalbumin (BLA) is studied by intrinsic fluorescence and circular dichroism (CD) spectroscopy. Both the nature and the kinetics of conformational changes of BLA upon adsorption on a polystyrene/water interface are characterized. Several theoretical models exist that describe the kinetics of adsorption as well as the kinetics of adsorption-induced conformational changes (2.Andrade J.D. Hlady V. Adv. Polym. Sci. 1986; 79: 1-63Crossref Google Scholar). The experimental difficulty of discriminating between both kinetic processes has hampered the experimental verification of these models. Here we show that the kinetics of adsorption and the kinetics of adsorption-induced conformational changes can be distinguished. In fact, it is demonstrated that the kinetics of both the diffusion and the adsorption processes do not need to be taken into account to enable analysis of the kinetics of the conformational change of the adsorbed protein as observed in stopped-flow experiments. Bovine α-lactalbumin (l-5385, Sigma Chemical Co.) was used without further purification. Polystyrene nanospheres, supplied as a colloidal suspension in water, were obtained from Polymer Laboratories (Heerlen, The Netherlands). Nanosphere suspensions were diluted with buffer before use. All experiments were done in a 10 mm Tris/HCl buffer of pH 7.50 containing 1 mmCaCl2. The added calcium ensures a constant calcium concentration during the adsorption and unfolding experiments, because calcium has a drastic effect on the stability and folding behavior of BLA. Nanopure water was used in all experiments (Sybron Barnstead NANOpure II). Dynamic light scattering was performed on an experimental setup composed of a Lexal 150-milliwatt laser operating at a wavelength of 514.5 nm, a photomultiplier, and a personal computer containing an ALV5000 correlation card. Scattered intensities were recorded at a 90° scattering angle and at 20 °C. A total of 10 measurements of each 10 s were averaged. The data were analyzed by the ALV5000 software. Samples were diluted in the buffer mentioned above. The nanosphere concentration ranged from 0.05 to 0.25 mg/ml. The electrophoretic mobility of the polystyrene nanospheres (3 mg/ml) suspended in a 10 mmTris/HCl buffer at pH 7.5 with 1 mm CaCl2 was determined by microelectrophoresis (Malvern Zetasizer III). The Malvern computer program was used to calculate the zeta-potential from the electrophoretic mobility. The adsorption isotherm was determined according to the depletion method. The amount of free protein after adsorption was measured spectrophotometrically using a molar extinction coefficient of 28,540 m−1 cm−1for BLA at 280 nm. The polystyrene nanospheres were separated from free α-lactalbumin by using Centricon 100 ultrafiltration devices (Millipore Corp.). Protein adsorption on the filter was checked and found to be negligible. The filter retained more than 99.8% of the polystyrene nanospheres. In a typical adsorption experiment 2.5 ml of protein solution was added to 2.5 ml of nanosphere suspension in a 10-ml polysulfone centrifuge tube (Nalgene, Oak Ridge, TN). The tubes were then placed on a rocking table and gently shaken at room temperature. Adsorption isotherms were determined after 30 min of adsorption and after 24 h of adsorption. Different protein concentrations were used to construct an adsorption isotherm. The resulting adsorption isotherm was used to determine the molar protein-to-nanosphere ratio (n) that gave maximal protein monolayer adsorption and minimal free protein. This ratio was used in all adsorption experiments, unless mentioned otherwise. For determination of the desorbed amount of BLA, the nanospheres covered with BLA where separated from the solution with Centricon 100 ultrafiltration devices. Separation was done at 4 °C without calcium or at 20 °C and in the presence of either 10 or 100 mmCaCl2. Subsequently the concentration of BLA in solution was determined spectrophotometrically. Polystyrene nanospheres (80 nm) were incubated with different amounts of CaCl2 (0, 5, 10, 40, 100, 200, and 400 μm) for 20 h at room temperature in a calcium-free 10 mm Tris/HCl buffer at pH 7.5. The nanospheres covered with calcium were separated from the calcium containing solution by ultrafiltration (Amicon, YM50 membrane, Millipore Corp.). The filtrate was analyzed for calcium content by inductively coupled plasma-optical emission spectrometry (ICP-OES). The amount of adsorbed calcium was calculated from the original amount and the amount of free calcium after filtration. An adsorption isotherm was constructed and fitted with a Langmuir isotherm: [Ca]adsorbed = (B*[Ca]free)/(Kd + [Ca]free). In this equation, B is the plateau value of adsorption, and Kd is the dissociation constant. The concentration of CaCl2 stock solutions was determined with ICP-OES. Fluorescence spectra and time-dependent fluorescence traces were measured in a quartz cuvette (10 × 4 mm) on a Fluorolog 2 (SPEX). The temperature of the cuvette holder was maintained at 20 °C with a water bath in all experiments. Excitation was at 300 nm, with excitation slits at 3 nm and emission slits at 5 nm. A blank, containing all components except protein, was subtracted from each sample. Nanosphere concentrations ranged up to 0.05 mg/ml. The total absorbance of the samples at 300 nm was kept below 0.1 to minimize the inner filter effect. To ensure a molar protein to nanosphere ratio of 500, the maximum protein concentration was about 0.5 μm. In the manual mixing adsorption experiments the nanospheres were added to the stirred protein solution with an automatic pipette. Typically, an amount of 100 μl of nanosphere suspension was added to 1.5 ml of protein solution. In time-dependent fluorescence measurements, care was taken to avoid photobleaching of the fluorophore, by ensuring discontinuous illumination with enough (dark) time between data points. Fluorescence anisotropy was measured on a home-built fluorometer equipped with two photomultipliers arranged in T-format (Thorn EMI 9863QA/350, operating in photon-counting detection mode). The light was generated by a 150-watt short arc xenon lamp, and the excitation wavelength of 300 nm was selected in a monochromator (Bausch and Lomb) with a bandpass of 3.2 nm. Polarizers were used in both the excitation light path (rotatable Glan Taylor polarizer) and the emission light path (Polaroid, sheet). The emission light was selected with a 335-nm cut off filter and a UG1 filter (Schott). A blank measurement, containing all components except BLA, was subtracted from each sample, and five measurements were averaged for each sample. Stopped-flow fluorescence and stopped-flow fluorescence anisotropy were measured on a BioLogic SFM4 equipped with a 2- × 2-mm cuvette (FC-20). Excitation was at 300 nm, and excitation slits were set at 0.5 mm, resulting in a bandpass of 4 nm. A cut-off filter of 335 nm (335FG01–25, Andover Corp.) and a 330wb60 bandpass filter (XF3000–25, Omega Optical Inc.) were used together to select the emission light. The temperature was kept constant at 20 °C with a thermostatic water bath. Ten measurements were averaged for each sample. In a typical run 150 μl of a 5 μm protein solution was mixed with 150 μl of a 10 nm nanosphere suspension in 75 ms. A flow speed of 4 ml/s resulted in a dead time of 15 ms. Faster flow speeds were not used, because this resulted in signal distortion and bad reproducibility, probably due to cavitation. Results from the stopped-flow fluorescence experiments were analyzed and fitted with the Padé-Laplace algorithm in the Bio-Kine software (Bio-Logic), using a minimum number of exponential phases. Prior to fitting the data, the signal of the polystyrene nanospheres was subtracted from the sample, and the time axis was adjusted, resulting in a shift of the first data point to t = 0 s. A fit was evaluated and judged to be correct when random noise around a horizontal line centered at zero was observed for the residual values. Stopped-flow fluorescence anisotropy was measured with a single photomultiplier and no polarizer in the emission light path. A similar set up has been reported recently in a study on protein folding in solution (23.Canet D. Doering K. Dobson C.M. Dupont Y. Biophys. J. 2001; 80: 1996-2003Abstract Full Text Full Text PDF PubMed Scopus (41) Google Scholar). In the anisotropy set-up the excitation light is modulated with a photoelastic modulator to give alternating horizontally and vertically polarized light with a frequency of 100 kHz. The synchronized electronics connected to the photomultiplier simultaneously record the fluorescence emission intensity at vertical excitation (Iv) and at horizontal excitation (Ih). A blank, containing all components except BLA, was subtracted from the sample resulting in the corrected values Iv* and Ih*. The anisotropy (A) for each data point was then calculated in a spreadsheet application according to the equation: A = (Iv* − Ih*)/(Iv* + Ih*/2). CD measurements were performed on a Jasco J-715 spectropolarimeter, equipped with a Peltier temperature control system set at 20 °C. Calibration was performed with an ammonium d-10-camphorsulfonate solution in nanopure water for which the concentration was checked spectrophotometrically. Typical protein concentrations used were 1.5 μm in a 0.1-cm cuvette for far-UV CD and 4.5 μm in a 1-cm cuvette for near-UV CD measurements. Eight scans were averaged for each sample. The CD spectra of BLA adsorbed on the nanospheres were averaged from 160 and 128 scans for the far-UV region and near-UV region, respectively. In the case of adsorbed BLA, sample and blank spectra were recorded alternately in sets of 32 scans, to avoid disturbances due to instrumental drift. For each set of 32 scans, a fresh sample of BLA adsorbed on the nanospheres was used. The response time was 1 s, and the spectral bandwidth was 1.0 nm. Before analysis of the spectra, a blank, containing all components except BLA, was subtracted from the sample. The radius of the nanospheres diluted in a 10 mm Tris/HCl buffer at pH 7.50 with 1 mm CaCl2 is 47 ± 1 nm as determined by dynamic light scattering. Based on this value a surface area of 122 ± 3 m2/g is calculated. Investigation by transmission electron microscopy shows that the nanospheres are spherical and that possible irregularities at the surface are smaller than 2 nm in size (results not shown). The zeta-potential for the nanospheres as determined by electrophoretic mobility is −38 ± 1 mV in a 10 mm Tris/HCl buffer with 1 mmCaCl2 at pH 7.5. This value indicates a negative surface charge of the polystyrene nanospheres in this particular buffer. The diameter of the nanospheres after adsorption of a monolayer of BLA is 49 ± 1 nm and remains the same during 24 h. No aggregation of the nanospheres occurs after adsorption of BLA. The adsorption isotherm of BLA on polystyrene nanospheres is shown in Fig. 1. The plateau value of 3 ± 0.2 mg/m2 agrees well with previously reported values. Adsorption of BLA on a platinum surface resulted in a value of 2.9 mg/m2 (24.Cabilio N.R. Omanovic S. Roscoe S.G. Langmuir. 2000; 16: 8480-8488Crossref Scopus (40) Google Scholar), and a value of 3.3 mg/m2 was found for adsorption of BLA on a hydrophobic silicon surface (25.Suttiprasit P. Krisdhasima V. McGuire J. J. Colloid Interface Sci. 1992; 154: 316-326Crossref Scopus (137) Google Scholar). The steep part in the region of low BLA concentration is an indication for high affinity behavior. In the initial, steep part of the adsorption isotherm, all protein present in the system is adsorbed on the surface of the polystyrene nanospheres and virtually no free BLA is present in solution. If a molar nanosphere-to-protein ratio of 1:500 is chosen, then all BLA added is adsorbed and there is no free BLA present in the solution. This ratio corresponds with the region in the adsorption isotherm of the initial steep part, before a coverage of 3 mg/m2 is reached. This ratio is used in the adsorption experiments presented below, assuming that all added protein is adsorbed on the nanospheres. The polystyrene nanospheres and the BLA molecules both have a net negative charge under the conditions used. The fact that adsorption occurs means that the electrostatic repulsion has to be overcome. Hydrophobic interactions therefore contribute the most to the adsorption process, as was found for the adsorption of BLA on hydrophobic interfaces (15.Haynes C.A. Sliwinsky E. Norde W. J. Colloid Interface Sci. 1994; 164: 394-409Crossref Scopus (156) Google Scholar, 26.Larsericsdotter H. Oscarsson S. Buijs J. J. Colloid Interface Sci. 2001; 237: 98-103Crossref PubMed Scopus (98) Google Scholar). However, minor contributions of electrostatic interactions cannot be excluded. They may play a role in the orientation of the adsorbed BLA molecules (see "Discussion"). Desorption of BLA from the polystyrene nanospheres is tested with different methods. Washing the BLA-covered nanospheres with buffer does not result in protein desorption, even after 24 h. Hardly any desorption of BLA from the polystyrene nanospheres takes place when the calcium concentration is increased. Only 12% of the adsorbed BLA molecules is desorbed when the calcium concentration is increased to 10 mm, whereas only 14% is desorbed at a calcium concentration of 100 mm after 24 h of incubation. In an attempt to decrease the hydrophobic interactions and enable BLA desorption the temperature was decreased. However, storage during 24 h at 4 °C and in the presence of 1 mmCaCl2 gave no desorption of BLA molecules. Fluorescence spectra of BLA in solution and of BLA adsorbed on nanospheres after 5 and 75 min of adsorption are shown in Fig. 2. The spectrum of BLA in solution shows a fluorescence emission maximum at 325 nm, which indicates that the Trp residues are buried in the hydrophobic interior of the protein. The adsorbed BLA shows a maximum at 335 nm and an intensity increase at maximum wavelength of about 100% compared with the spectrum of BLA in solution. The shift of the fluorescence maximum indicates that the tryptophan residues of BLA in the adsorbed state are in a more exposed environment. The intensity increase is related to a reduction in quenching that results from a different position of the tryptophan residues compared with their original position in the three-dimensional structure of native BLA. For native BLA it is suggested that energy transfer occurs from Trp26 and Trp104 to Trp60 and, furthermore, the fluorescence is quenched by two disulfide bonds near Trp60(Cys73-Cys91 and Cys61-Cys77) (27.Arai M. Kuwajima K. Fold. Des. 1996; 1: 275-287Abstract Full Text Full Text PDF PubMed Scopus (158) Google Scholar). It has also been observed that unfolding of BLA results in a red shift of the fluorescence emission maximum and an accompanied increase in fluorescence intensity (28.Ostrovsky A.V. Kalinichenko L.P. Emelyanenko V.I. Klimanov A.V. Permyakov E.A. Biophys. Chem. 1988; 30: 105-112Crossref PubMed Scopus (28) Google Scholar). Complete unfolding of α-lactalbumin by 5.4 mguanidinium hydrochloride results in a fluorescence emission maximum of 350 nm indicating complete exposure of the Trp residues (results not shown). A measured fluorescence emission maximum of 335 nm would thus indicate that the adsorption-induced unfolding is not complete, leaving one or more Trp residues in the hydrophobic core of BLA. Circular dichroism spectra of free and of adsorbed BLA in the far-UV region and in the near-UV region are shown in Figs.3 (A and B, respectively). The far-UV CD spectrum of adsorbed BLA shows that a considerable amount of secondary structure remains after adsorption. Although the spectrum of adsorbed BLA contains a lot of noise, the characteristic ellipticity minimum at 208 nm is visible. This indicates that the secondary structure of native BLA and of adsorbed BLA is similar. The near-UV CD spectra show a clear difference between native BLA and adsorbed BLA. The spectrum of BLA in solution has a characteristic profile that includes a broad Tyr minimum around 270 nm and a local Trp minimum at 297 nm. The spectrum of adsorbed BLA does not show these characteristics and has a decreased molar ellipticity over the entire near-UV CD spectrum, which indicates disruption of tertiary structure elements. The latter is in agreement with the changes seen in the fluorescence emission spectrum upon protein adsorption (see above). The difficulty of working with polystyrene nanospheres in suspensions emerges in particular when working with far-UV light in CD spectroscopy (Fig. 3A). The CD measurements in the far-UV region were performed with a nanosphere concentration that results in a UV absorption of 0.5 at 220 nm in a 0.1-cm light path cuvette, whereas in the near-UV a concentration of nanospheres is used, which results in a UV absorption of 0.5 at 280 nm in a 1-cm light path cuvette. The protein concentration was adjusted to obtain a ratio of 500 protein molecules to 1 nanosphere particle, as is used throughout this work. The loss of light intensity in the CD measurements of the adsorbed protein, which is due to both light scattering of the polystyrene nanospheres and light absorption of the polystyrene molecules in the nanospheres, leaves only half of the original intensity for the characterization of the protein structure. Consequently, a significant increase of noise in the data is observed. Despite the difference in the noise level between the spectra shown in Fig. 3, it is clear that upon BLA adsorption a disruption of tertiary structure elements occurs without significant loss of secondary structure. Stopped-flow fluorescence spectroscopy is used to investigate the kinetics of the adsorption process. To our knowledge, we demonstrate here for the first time protein adsorption on solid spheres in the stopped-flow technique. The small size of the nanospheres allows the use of this technique, without the risk of blocking the small mixing channels in a stopped-flow apparatus. A typical example of a stopped-flow trace is shown in Fig. 4. The experiments are performed with different BLA/nanosphere ratios ranging from about 1100 to 160 BLA molecules for each nanosphere. This ratio covers the steep part of the adsorption isotherm as descri
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