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Application of fluorescence spectroscopy to the study of proteins at interfaces
Application of Fluorescence Spectroscopy to the Study of Proteins At Interfaces ALAN G. WALTON AND FRANK C. MAENPA a Department o f Macromolecular Science, Case Western Reserve University, Cleveland, Ohio 44106 R e c e i v e d N o v e m b e r 16, 1978; accepted M a r c h 16, 1979 A n e w technique for studying the nature of adsorbed protein molecules b a s e d on fluorescence s p e c t r o s c o p y is described. T h e m e t h o d was u s e d for following the adsorption of bovine s e r u m albumin (BSA) onto particulates. On r a n d o m copolypeptide substrates of the type (Glu(OBzl)XAlaU), (Glu(OBzl)XLeu'), or (Lys(CBZ)XLeu,), considerable fluorescence q u e n c h i n g was observed. The origin o f this effect appears to be a combination of the energy e x c h a n g e b e t w e e n the excited t r y p t o p h a n residue of B S A and the excited triplet state (*Ba2b) or of 1Lo singlet state of the substrate, a n d q u e n c h i n g by adjacent molecules. Since energy e x c h a n g e o f this type can o c c u r over a distance of 100 ]k or so, it is possible to probe the protein e n v i r o n m e n t of particles in the interracial layer. It is c o n c l u d e d that e v e n at low surface coverage, protein molecules are p r e s e n t adjacent to the surface in a n o n a d s o r b e d " m e t h o r i c " layer.
INTRODUCTION
The interactions between proteins and surfaces are of great importance in nature, probably beginning in the prebiotic stages of chemical evolution, where rock surfaces may have played a role in polymerizing and organizing primitive proteins or proteinoids (1). In the most sophisticated biological structures present today, proteins or glycoproteins play an important role at interfaces including cells, joints, and arteries. Indeed, it is widely believed that protein adsorption at the blood vessel wall plays an integral role in control or activation of thrombogenesis (2). Questions concerning the behavior of proteins at "foreign" or nonbiological surfaces have been brought forth by the need to develop blood-compatible materials and prostheses (3) but are also important in various forms of chromatography, and in all cases where animal or vegetable matter 1 Current address: U. S. Biochemical Corp., Warrenville H t s . , Ohio.
come into contact with plastics or other surfaces. Following insertion of a polymer or other material into blood, proteins, particularly serum albumin and fibrinogen, are adsorbed fairly rapidly. Blood clotting can be initiated by the activation of one or more blood clotting factors (proteins) on interaction with the surface and/or by stimulated release of proteins and other entities from blood cells as they interact with the surface. In view of the central importance of protein behavior at interfaces, considerable effort has gone into characterizing the process (4). Approaches to the study of proteins at interfaces have considered two main facets of the problem--how much and what type of protein is adsorbed and what, if anything, happens to the structure of the protein in the adsorbed state. The first problem may be approached conveniently by studying adsorption isotherms of one or more proteins on a welldefined substrate. Experimentally one is faced with the familiar problem in surface
265
Journal of Colloidand Interface Science, Vol. 72, No. 2, November 1979
0021-9797/79/140265-14502.00/0 Copyright © 1979by AcademicPress, Inc. All rights of reproduction in any form reserved.
266
WALTON AND MAENPA
chemistry of either providing a large enough surface area that adsorbs measurable amounts of protein, or finding a sensitive method or tag (usually radioactive labeling) that allows study of protein uptake on relatively small surface areas (of, say, polymer film). In an example of the former method Norde (5), Lyklema (6), and others have measured the uptake of serum albumin on polystyrene latex spheres by calorimetry. They find that an essentially irreversible uptake occurs, first to approximately monolayer coverage, and then beyond. They were unable to reach a definitive conclusion concerning structural changes of adsorbed protein. However, Nyilas (7), using albumin and a disperse silica substrate and calorimetry, concluded that adsorption from solution caused denaturation of the protein in the interfacial region. Kim and co-workers (8, 9) have measured the uptake of albumin and fibrinogen both singly and in competition on polymer films by labeling the protein with iodine. They find that the protein adsorbs to a monolayer and beyond but, in mixtures, the relative amount of each protein adsorbed is a function of the time of measurement and the nature of the substrate. Although radiolabeling of proteins provides a convenient method of measurement, there seems to be some concern that such a procedure can in itself cause minor changes in protein conformation a n d " stickine s s" (10). Another method of detecting the extent of protein adsorption at a flat interface has been ellipsometry (11). By this technique the extent of protein attachment is measured by assessing the thickness of the refracting layer. The technique is only well suited to smooth surfaces and particularly reflective surfaces and provides, at best, very limited information concerning the structure of adsorbed protein. Infrared differential spectroscopy has been used by Morrissey and Stromberg (12) to examine the attachment and conformation of bovine serum albumin (BSA) on silica. Conformational changes are difJournal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
ficult to assess for globular proteins in aqueous solution, but they concluded that no significant conformational changes occurred, even for low adsorbed concentrations in this system. In an attempt to provide more direct evidence of the conformation of adsorbed proteins on surfaces, we have previously presented a surface circular dichroic (C.D.) method in which the structure of a monolayer, or less, of an adsorbed protein was readily assessable (13). It was found that fibrinogen underwent no detectable conformational change for adsorption onto quartz, that there were slight changes with bovine serum albumin (14), and that there were considerable changes for the clotting activation protein, factor XII (Hageman factor) on the same substrate (15, 16). Whereas the surface C.D. method provides direct information on surface-adsorbed protein structure, it cannot be used if the substrate absorbs UV light strongly, is rough, is optically active, or cannot be cast in a thin film on a transparent substrate. To circumvent some of these difficulties, we have been exploring the application of fluorescence spectroscopy (17, 18). The potential advantages of such a technique are that it is very sensitive and has been used for determining environmental and structural changes in proteins, including those in cell membranes (19). In addition, since fluorescence occurs for most proteins in the 340-mm region and virtually no potentially useful substrate fluoresces at this wavelength, interference from the substrate is minimized. If the substrate absorbs ultraviolet light in the region of protein stimulation (280 nm) or emission, this effect can be corrected for. The sole disadvantage seems to be that, unlike C.D. spectroscopy, the data cannot be interpreted in terms of detailed structural changes. There are basically two approaches available for examining the fluorescence of adsorbed proteins. The first, expounded here, uses a particulate substrate and meas-
STUDY OF PROTEINS AT INTERFACES ures the amount of fluorescence quenching, i.e., the decrease in fluorescence during adsorption. This method uses the intrinsic fluorescence of the proteins which involves the fluorescence of tryptophan and sometimes tyrosyl residues in the protein chain. The second method proposed recently (20), involves internal reflectance fluorescence spectroscopy (IRFS) and can be applied to thin films coated upon reflectance crystals. This method involves measurement of fluorescence increase in the interfacial area but generally requires an extrinsic fluorophore. In the cited work (20), the adsorption of fluorescein isothiocyanate (FITC)-labeled y-globulin on silicone rubber films was measured. Each technique has advantages and disadvantages. In general it would be preferable to use a technique which relies on intrinsic fluorescence of proteins on limited-area films of reasonable thickness (>10 /~m). Neither method is capable of this resolution and indeed the electronic principles involved are different. The following report outlines some principles and experimental data for the adsorption of bovine serum albumin on particulate synthetic copolypeptide substrates. These random copolypeptides are similar to petroleum-based polymers in many of their properties and typically are based on L-glutamic acid and L-lysine as the acidic or basic components and L-alanine, L-leucine, and glutamic acid or lysine derivatives as the hydrophobic component. The structure of some of these units in a representation of part of the polymer chain is shown below.
T
-NH-C- CONH-C-CONH-
I (~H2)2
I CH3 L- olony[ residue
-eorbobenzoxy L-lysyl residue
|COOCH2 O 7 -benzyI-L-glufomyl residue
-NH-C-CONH-C-CONH-
] CH3
I CH3
NH
I
CBZ L-leueyl residue
267
This type of substrate was used because of its potential viability as a biomaterial (21, 22). Since polypeptides are in some ways similar to native protein materials, this type of substrate may act as a model biological interface. However, from an experimental point of view, the polypeptide could present particular difficulties since we seek to examine the adsorption (and conformation) of proteins consisting of a polypeptide chain, on a similar substrate. There has been a previous report from our laboratory of the adsorption of Hageman factor onto polypeptide substrates (23) in which it was shown that protein fluorescence is indeed modified by the substrate. The nature of the modification is explored in this work. EXPERIMENTAL The bovine serum albumin used in this work was Miles primary standard monomer. Copolypeptides for substrates were provided by Drs. J. M. Anderson and T. Hayashi and have viscosity molecular weights of 50,000 or more. Stock suspensions of amino acid copolymers were prepared by first dissolving 1 to 2 mg of polymer in about 2 ml of chloroform. A few drops of the resulting solution were shaken with about 50 ml of distilled water. The chloroform was driven off by gentle heating in a hot water bath with the aid of moderate evacuation by an aspirator. Care was taken to keep the chloroform emulsified by continuous shaking during the process. Once all the chloroform was driven off, the process was repeated until all of the polymer solution was used. The resulting aqueous suspension was kept under an aspirator vacuum for 1 hr to pull off the last traces of chloroform. The resulting suspensions were filtered through a coarse glass filter to remove any large particles that may have been present. Particulate suspensions were examined in a Coulter counter and an electron microscope (Hitachi Hu 11A). The particles were Journal o f Colloid and Interface Science, Vol. 72, No. 2, November 1979
268
WALTON AND MAENPA
20~
10C 0 Z
1'o
b
20 n r 2 x 5000,mm 2
4O
30
% t: 1000 Q.
u~ SO0
0
o
1'o
2'o
30
40
n r 2 x 5000, mm 2
FIG. 1. Histograms of particle surface area for polypeptide suspensions of Glu(OBzl)Z°:Leu 8° calculated from electron micrographs. (a) Histogram of particle surface area as a function of size for
Glu(OBzl)2°Leu8°. (b) Histogram of fraction of total surface area of Glu:Leu 1:4 vs particle surface area. Although there are a large number of smaller particles, their contribution to the total surface area is limited. essentially spherical but heterodisperse in size. Typical histograms are shown in Fig. 1, which represents the relative n u m b e r of particles that have a given surface area (estimated assuming a spherical shape), and the relative contribution to total surface area b y particles of a single size. The histograms were prepared from micrographs of 16 random segments o f the prepared grids. The particles showed the same X-ray diffraction and infrared spectrum as the bulk material (a-helix) and therefore have substantially the same structure as the bulk solid. Relative fluorescence intensity measureJournal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
ments were made using an A m i n c o - B o w man spectrofluorimeter with an Ortec photon counter. Adsorption measurements were made by first measuring the fluorescence of the protein solution in 0.001 M phosphate buffer at p H 7.0 and 25°C. The fluorescence of the particles in protein (in buffer) was measured on a separate sample (to avoid fluorescence bleaching) and the fluorescence of the remaining protein was measured from a third, equivalent sample after centrifuging. Prior to the actual adsorption experiments, a determination was made on the most effective centrifugation rates and times by using suspension blanks. The effective-
STUDY
OF PROTEINS
ness of the centrifugation parameters on separating the particles was evaluated by measuring the loss of scattered light intensity (in the spectrophotofluorimeter) of the centrifuged samples compared with a distilled water blank. Typically, less than 1% of the original suspension light scattered, and ultraviolet light absorbance remained after 2 hr of centrifuging at 20,000-30,000g. In these circumstances and at 4°C, no detectable amount of protein was removed from solution. Particles, with their adsorbed protein, were removed from protein solution by using the centrifugation method as described. Beam Attenuation Correction
Since the particles of the suspension are insoluble polypeptide derivatives, they adsorb and scatter ultraviolet light in the region of fluorescence excitation (282 rim) and emission (345 nm). Consequently the relative fluorescence intensity of the protein in the absence and presence of suspension would be different, even if no adsorption or other solution modification were achieved. This effect, beam attenuation, can be accounted for by measuring the ultraviolet spectrum at the two wavelengths, and introducing a correction factor. If, for example, the absorbance and scattering at 282 nm are 10% of the incident light intensity for a certain suspension in a 1 cm cell, then, because of the 90° scattering geometry of the fluorescence system, the average protein molecule in the fluorescence beam receives 5% less light. Similarly, if the absorbance and scattering at 345 nm are 10% of the emitted light, the correction is 5% based on the geometry of the instrument. Another way of explaining this correction is that light passes through only one-half of the cell until it is absorbed, and one-half (at right angles) after emission. The relation between fluorescence intensity and ultraviolet absorbance is discussed separately; however, we find that the loss of fluorescence intensity with
AT INTERFACES
269
various suspensions used here lies between 1 and 5%. Nature of BSA Fluorescence and Structure
Albumin is the most abundant protein in serum. Two slightly different albumins are in common use: human serum albumin (HSA), which consists of 585 amino acid residues; and bovine serum albumin, consisting of 581 residues, many of which are homologous with HSA. The primary sequences of both have been established (24, 25). There are two classes of fluorescing proteins, class A proteins that contain no tryptophan and class B containing one or more tryptophan residues. This distinction is made because, of the three amino acid residues that fluoresce, phenylalanine, tyrosine, and tryptophan, the first two are quenched in the presence of tryptophan. Thus for class A proteins the detectable fluorescence is due to tyrosine residues (the quantum yield of phenylalanine is very low) and class B protein fluorescence originates in tryptophan residues. Human serum albumin contains only one tryptophan residue (residue 214) and appears to be unique in that both tryptophan and tyrosine fluorescence are detectable. On the other hand, bovine serum albumin has two tryptophan residues (residues 134 and 212) and is strictly a class B protein (see Table I). On denaturation (i.e., disruption of the secondary and tertiary structure by heating) the fluorescence maximum shifts and decreases. For HSA it moves from 339 nm to 350 nm with a drop in quantum yield from 7.4 to 5.2 and for BSA there is a shift from 342 nm to 350 nm and a concomitant drop in fluorescence from 15.2 to 7.4. Clearly BSA fluorescence is more sensitive to conformational change (denaturation) than HSA and it is BSA that is used in this work. The BSA molecule is estimated to be 116 x 27 x 27 A from Langmuir trough measurements. The detailed three-dimensional Journal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
270
WALTON AND MAENPA TABLE I Comparison of Amino Acid Sequence in the Region of Tryptophan in Bovine and Human Serum Albumin BSA Asp Glu P-he] Lys Ala Asp 125
i
t
1
~
, kys kys iPhe~
Gly
I
130 I
~
= 135
*
140
HSA Thr A~, ffh_ej,i~ Asp A~nLG!,:Glu Th~'Zh_e:Leu Ly~',Syj_Tx~_L_~u_TL~_Le~_TL~_G1LUe 205
BSA
210
212
215
220
-Gfu-~u" "~rg-~a~ Leu rL}~--~a-[~, Set ~Va7-ATa-AFg-L~ -fer-gin'], Lys [Phe- P-to -ky~I
I
' : HSA .GIu . .GIu. Arg . . Ala . I. Phe . . Lys . .Ala. .
L
I
I
I
, , , . A1a . . iVal . . Ala . . Arg . . Leu . .Ser. .Glnl . Arg . . iPhe . . Pro . . Lys . .
structure of the albumins is not known; however, circular dichroism spectra indicate that >50% of the structure is a-helix in each case. There is internal homology in the structure, indicating that it probably consists of three equivalent structural units. The most unusual feature of serum albumin is the unique sets of C y s - C y s groups. These occur at residue numbers 90, 91; 168,169; 245, 246; 278, 279; 360, 361; 437, 438; 476, 477; and 558, 559. It is possible to perform a C h o u - F a s m a n type of conformational analysis (26) which shows, as expected, long
.
a-helical regions. Typically the sequence contains a large number of runs of the type -P-N-N-N-P-N-N-P-, where P is a polar residue (Arg, Lys, Glu, Asp) and N is nonpolar. Such sequences are typical of a-helices in which the - P - residues lie in contact with the solvent (water) and the remaining residues form a hydrophobic interior. With these features in mind it is possible to postulate a crude structure in two and three dimensions. Figure 2 shows a model for human serum albumin which is essentially a set of a-helices of length 72-82 residues (110-120 ~i) linked
Hyd rophobic interior
- - ~- helices
o
cys-cys
120A
~80 residues long
pairs
:::X
Irregular region
30A
,,
'
FIG. 2. Schematic representation of serum albumin, based on conformational analysis and known geometric properties. Journal of Colloid and Interface Science,
Vol. 72, N o . 2, N o v e m b e r 1979
STUDY
OF PROTEINS
271
AT INTERFACES
0.8
0.6
G)
0.4
O e., .O
ol e~ ,~ 0.2
i
2
4
6
B.S.A.
8
10
(//g/ml)
FIG. 3. Relative fluorescence intensity and absorbance of BSA in /zg/ml range. The line C corresponds to measured values; A is corrected for the exponential term in Eq. [3]. by di-Cys bridges. The hydrophobic N and C termini are imagined to form a hydrophobic lock down one side of the cylinder and residues 246-278 and 438-476 form a base of irregular structure. On this basis one has 3 8 6 - a - h e l i x bends/585 = - 6 5 % ahelix. The structure has a hydrophobic " h o l e " down the middle, capable of carrying fats or other hydrophobic material (one of albumin's main functions). The Trp residues in H S A and BSA lie in regions that are probably o~-helical. The model will serve as a reference but should not be taken too literally until substantiated by further evidence. Fluorescence occurs when an electronic excited state cannot dissipate its energy through excited vibrational and/or rotational states, i.e., by thermal decay. Indole ring systems, of which tryptophan is an example, fall into this category. Absorptive transitions o c c u r between the ground state (A) and first or second excited state 1La or 1Lb. It is construed that decay occurs by intersystem crossing to the 3La triplet state with fluorescence occurring by decay of that state to a vibrationally excited state (27).
The fluorescence observed at a constant exciting-light intensity is equal to the light absorbed multiplied by a factor q5 which is dependent upon the quantum yield and an instrumental factor: Fx -
6(lo - I)x
,
[1]
Io where Io is the incident and I the radiated or transmitted intensity. These factors may be related by the B e e r - L a m b e r t equation
axLc = 2.303 loga0 (Io/I)x,
[2]
where eX is the molar extinction coefficient, L the path length, and c the concentration. Then Fx = ~b(1 - e x p [ - e a c L ] ) .
[3]
Thus for small absorbance the B e e r - L a m bert relation should hold with fluorescence replacing absorbance. Figure 3 shows that Eq. [1] holds for BSA up to a concentration of - 5 /zg/ml; the method is accurate to 0.1 /zg/ml or better, and thus provides a useful concentration assay. Journal of Colloid and Interface Science, Vol. 72, No, 2, November 1979
272
WALTON AND MAENPA 100
342 nm is quenched by the presence of (Glu(OBzl))8°Ala 2° copolypeptide particles. There is approximately a 25% reduction of fluorescence caused by the particles, but there is no indication of a wavelength shift to 350 nm that would be indicative of protein denaturation. The effect is similar to the previously noted decrease of Hageman factor fluorescence on polypeptide particles (23). Using the correction procedure given in the experimental section, it is possible to show that 1-5% of the reduction in fluorescence is caused by absorbance and/or by the polypeptide particle scattering. The remainder is due to a genuine loss of fluorescence of BSA protein in the presence of the glutamate copolymer. The decrease in fluorescence, AF, increases with increasing protein concentration and substrate particle density and is clearly identifiable with a surface interaction. In order to establish what fraction of the surface adsorbed protein was fluorescing, the particles were removed from solution by careful centrifugation, avoiding centrifugation of solution protein, as described in the experimental section. By this means, the loss in protein from solution could be ascribed to surface adsorption. Figure 5
8O
60 _= c
u.
~ if'
h 20
:l..'l
•'7 I
200
..... i
I
I
300
I
400
soo
~./nrn
F ~ . 4. F l u o r e s c e n c e of B S A ( ) in the a b s e n c e of (Glu(OBzl)8°:Ala2°); ( - - - ) in the p r e s e n c e of particles of the copolypeptide; ( . . . ) after r e m o v a l of copolypeptide particles with a d s o r b e d protein. RESULTS
Figure 4 shows the result of a typical run in which the fluorescence of BSA at
1.0
GL
1:4
.8
o
e~ <~'°
•
x
O
1= .4 xo
I--
,2 BSA
.4 in soln., mg/m I
.6
.8
1~0
FIG. 5. Typical adsorption i s o t h e r m s for B S A adsorbed on (Glu(OBzl)2°:Leu 8°) copolypeptide particles in the low coverage ( < 10%) m o n o l a y e r region b a s e d on fluorescence and electron m i c r o s c o p e data. (Symbols × a n d O refer to separate runs.) Journal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
273
STUDY OF PROTEINS AT INTERFACES T A B L E II
surface remodeling, aggregation, and perhaps surface poisoning by random entities. However, the most startling feature is that, if the surface-adsorbed protein is fluorescing, the total fluorescence should decrease after centrifugation. In fact it increases as indicated in Fig. 4. This discrepancy cannot be explained by particle absorbance of light and/or scattering, both of which have been measured carefully. For some reason (that will become apparent shortly) protein adsorbed on the surface and in the immediate area of benzyl-group-blocked polypeptide substrates, does not fluoresce. We have compared these experiments with particles of (Glu2°:Alas°) copolymer and with colloidal silica. In neither case is there a decrease in fluorescence even though protein is adsorbed. Consequently particles of copolymers Glu(OBzl):Leu, Glu(OBzl): Ala, and Lys(CBZ):Leu cause fluorescence quenching of BSA, whereas the Glu:Ala polypeptide and silica do not cause such a change.
Table of B S A A d s o r b e d on Copoly-(Glu[OBzl]8°:Ala 2°) Showing Strong and W e a k A d s o r p t i o n L a y e r s a
Total BSA /~g/rnl
Adsorbed BSA (calc) (/xg/ml)
23.75 95 237 713
9.4 23.0 46.8 113
Adsorbed BSA (cent)° (/zg/rnl)
BSA stripped~ (/zg/rnl)
7.0 16.2 20.5 54
2.4 6.8 26.3 59
a Equilibration with protein for 4 - 6 hr. b Cent. refers to material centrifuged with the particles. c On the a s s u m p t i o n that the stripped protein does not fluoresce at all in the p r e s e n c e of the polymer.
shows the low-coverage region in which the uptake of protein by surface is essentially a linear function of protein concentration. If the suspension is left to stand, the rate of protein uptake diminishes rapidly, dropping to about one-half in a period of 24 hr. We ascribe this feature to a combination of
0~ 300
0.~."0~0-
5mm cell
/
/ /
200
0
O
ofo ~
° ~ o ~
lOmm cell
100
/° I
I
lOOO
BSA
I
(/~glmg)
3000
FIG. 6. Relative fluorescence o f B S A in high-concentration region showing s e c o n d a r y scattering effects. Journal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
274
WALTON AND MAENPA
1l
.13
.12
"'e
IJ.
I: O N ~
.11
.10
g .09,
i
~,
BSA
mg/ml
FIG. 7. Polarization ratio of BSA as a function of concentration. This information indicates that association of molecules does not occur at these concentrations. The absolute accuracy of the measurements is in the range 10-15% and no significanceis attributed to the scatter at low concentration. If the difference between the fluorescence of the protein in the presence of particles (corrected for light absorbance and scattering) and the fluorescence of protein after removal of the particles is attributed to a 100% quenching of protein influenced by the substrate, it is possible to calculate this amount of protein. Table II compares the amount of protein adsorbed with an additional amount not fluorescing in the presence of particles. Of course if the protein influenced by, but not adsorbed to, the particles is only partially fluorescing, then the quantity of material involved must be increased accordingly. DISCUSSION
Searching for an explanation of the fluorescence behavior of BSA in the presence of benzyl-esterified polypeptide particles, there seem to be three possibilities. The first is that the particles cause denaturation of adsorbed protein. H o w e v e r , such a postulate does not explain the increase in fluorescence after centrifugation. Journal o f Colloid and Interface Science, Vol. 72, No. 2, November 1979
Second, it seems possible that when protein molecules are crowded together on the surface or in the surface boundary layer, they undergo self-quenching. Such an effect could originate either in multiple scatter of light or in an energy-transfer process. There is some evidence that such a process can occur: At concentrations of BSA (>1 mg/ml) the fluorescence of BSA is attenuated. If the cell path length is halved, the fluorescence increases, indicating mutual interference effects (see Fig. 6). Consequently if high concentrations of protein occur in the interfacial region some attenuation and intermolecular quenching may occur. Crowding together of molecules could be a result of molecular association in the form of dimers, trimers, etc. Such an association can be tested by assessing the fluorescence depolarization. If molecules b e c o m e associated, their rotational diffusion is slowed and hence the rotation between absorbance and emission is changed. If molecules are associating there is an increased fluorescence polarization with increase in concentration. Figure 7 shows that there is only a small polarization of BSA
275
STUDY OF PROTEINS AT INTERFACES
solutions which is independent of the protein concentration. Similarly the introduction of particles does not produce a detectable increase in polarization of fluorescence. The fact that the quenching effect is not seen with colloidal silica seems to indicate that neither of the above effects is the major contributing process. The third possibility is that of molecular fluorescence quenching. If there is an energy acceptor within range, it is possible for the tryptophan residues to lose electronic energy by energy transfer. The efficiency of energy transfer is given by E -
1 1 + (R/Ro) 6
,
[51
where R is the separation of donor and acceptor and R0 is a characteristic distance. Energy transfer generally is detectable in the rangeR = 10-100 A. Literature reports have concentrated exclusively upon the transfer of energy within the same molecule, for example, between Trp and e-N-dansyllysine in adrenocorticotropin hormone. The dansyl group acts as an excellent fluorescence acceptor (28). In the present case we propose that energy-transfer processes can occur between the Trp residue of BSA and the benzyl radical of the substrate. To our knowledge energy transfer between a protein and solid substrate has not been documented previously. In addition to the energy transfer with acceptor groups in the substrate, it is conceivable that energy transfer can occur between protein molecules in close proximity to one another, either on a surface or in an adjacent layer. According to the F6rster theory of energy transfer, the separation distance R is related to the molecular parameters by R =
K2 8.79 × 10--28JAD n 4
1 -- T E ]1/6
× 4 ~ ' ~
.
[6]
~ LySCBZ2O 0Leu8 ~"~~.._ a,.oBzrOA,a~O , !
I
200
300
t 400
,~- (nrn)
FIo. 8. Ultraviolet spectra of Glu(OBzl)8°:Ala2° and Lys(CBZ)2°:Leu s° showing the split 1Lo transition in the 258-nm region and absorbance in the 30-nm+ region.
TE is the transfer efficiency given by
TE=
KT Kr + KQ + KR
K~,, KF, and KQ are the first-order rate constants for nonradiative transfer of excitation energy, fluorescence emission, and internal quenching of radiation, respectively. JAD is an "overlap integral" which accounts for the overlap between absorbance and fluorescence spectra of the acceptor. The quantity 4~°ois the fluorescence quantum yield of the donor without energy transfer and K / n ~ is a factor accounting for the relative orientation of donor and acceptor. Of the preceding factors it is readily possible and important to establish the overlap integral JAD. If the fluorescence spectrum of Trp and the absorbance spectrum of the Journal of Colloid and Interface Science, Vol. 72, N o . 2, N o v e m b e r 1979
276
WALTON AND MAENPA 80
Glu(OBzl):Glu does not associate with BSA. Since the average separation of molecules at the concentrations used is > 1000/~, there is no effective energy transfer. On the other hand, the polycation polylysine interacts with the polyanion B SA; the Lys(CBZ): Lys copolymer should also complex. Figure 9 shows the fluorescence spectrum of the basic copolymer with BSA; clearly the fluorescence is heavily quenched. A study of the conformation of the components BSA and poly-L-lysine by difference C.D. spectroscopy shows no net change of conformation on mixing (see Fig. 10). There is, therefore, no concurrent denaturation of the protein that could account for fluorescence quenching.
6O ta e-
=
-/ c
,~ 40
•~
2o
~,
/
I
250
I
350
I
450
M.m FIG. 9. Fluorescence spectra of water-soluble polypeptides (a) Glu(OBzl)6°:Glu4° and BSA, and (b) Lys(CBZ)5°:Lys5° and BSA (1:1 weight mixtures in each case). substrate do not overlap, then JAD ---- 0 and no energy transfer can occur. Figure 8 shows the absorbance spectrum of Glu(OBzl)S°Ala 2° and Lys(CBZ)2°:Leu s° in the 300-nm region. To benzene and benzyl groups are generally ascribed three major bands in the 185- to 195-nm (aBa), 205- to 225-nm (1La), and 255- to 275-nm (1Lb) regions.The 1Lb band of the two copolypeptides centered at 258 nm slightly overlaps the Trp excitation band in the 300-nm region. H o w e v e r , there is a forbidden transition ~Ba,b for benzene in the 360-nm region (not seen in the polypeptide spectra) that has almost complete overlap with the BSA fluorescence spectrum. In order to test whether energy transfer is indeed the predominant mechanism of energy transfer, we h a v e prepared modified samples of Glu(OBzl)~°°-x:Glu x and Lys(CBZ)l°°-ZLys x that are water soluble. In solution we look for fluorescence quenching caused by the association of the polypeptides with BSA. T h e acidic p o l y m e r Journal of ColloM and Interface Science, V o l . 7 2 , N o . 2, N o v e m b e r
1979
I.U ,.,I
o c&d
a E 0
o U,I a
200
210
220
230
WAVELENGTH
240
250
(nm}
FIG. 10. Circular dichroism spectra of (a) BSA (0.1 mg/ml); (b) poly-L-lysine (1 mg/ml); (c) addition of spectra (a) and (b); (d) spectrum produced by mixture of poly-L-lysine and BSA. Curve (a) has been reduced in scale for purposes of presentation. (The eUipticity of BSA at 220 nm is -18,000 deg.cm2/ dmole; polylysine at 220 nm is + 5000 deg. cm2/dmole.)
STUDY OF PROTEINS AT INTERFACES
277
We conclude, therefore, that the origin not scatter light unduly or absorb ultraof fluorescence quenching of BSA on poly- violet light in the 300- to 600-nm region is peptide particles lies predominantly in required. Suitable substrates include incharge transfer from the molecules adhering organics (glass, etc.) or organics low in to the surface, and those in the interfacial double bonds and devoid of benzyl or conregion (particularly within a distance of jugated benzene groups. In the case where 100 A). It seems likely that if the molecule the substrate does act as an energy-transfer lies flat on the surface the fluorescence will agent, some idea of the surface environbe heavily quenched since both tryptophan ment can be obtained by determining deresidues will be in close proximity to the tails of quenching processes. Polymers substrate acceptor. On the other hand, per- expected to act as energy-transfer systems pendicular adsorption is likely to cause a would include polystyrene, polyphenyl, etc. greater separation of Trp and substrate, with decreased energy transfer. ACKNOWLEDGMENTS The loose layer, which is swept free on We wish to acknowledge the experimental assistance centrifugation, could consist of perpendic- of Mr. Mark Soderquist and Dr. K. Raina. The ular molecules and others in a surrounding polypeptides were supplied by Dr. James Anderson "methoric" layerY In any case the Trp at Case Western Reserve University and Dr. T. moieties in these molecules must lie within Hayashi of Kyoto University. Modified polymers were 100 A of the surface for quenching to occur. produced by Dr. J. Caplin. This work was carried CONCLUSIONS
The fluorescence data point to the formation of a strongly adsorbed set of BSA molecules that probably are flat on the surface. Other molecules are less strongly adsorbed and form a "methoric" layer. The concentration of loosely attached molecules is at least as high as that of strongly attached molecules. The mechanism of energy transfer is probably via a combination of the triplet (3Ba,o) and 1Lb states of the substrate and interaction with adjacent bound or unbound molecules. No evidence has been found of denaturation of BSA on adsorption on these substrates (though this is not excluded). We conclude that fluorescence measurements can only determine directly whether conformation changes accompany adsorption when there is no mechanism for energy transfer with the substrate. A particulate suspension of substrate that does 2 The term "methoric layer" has been widely used in the colloid literature by the Yugoslav school to mean dense surrounding layers of counter'ions. In this case we use the term to indicate dense interfacial layers of protein.
out under the auspices of a Program Project in Biological Materials N 1HLB 15195. The junior author (F.C.M.) was supported by an N.I.H. Traineeship Award. REFERENCES
1. FOX, S. W., and Dose, K., "Molecular Evolution and the Origin of Life," p. 196. Freeman, San Francisco, 1972. 2. Salzman, E. W., Blood 38, 509 (1971). 3. Brash, J. L., and Lyman, D. J., in "Chemistry of Biosurfaces" (M. L. Hair, Ed.), p. 177. Dekker, New York, 1971. 4. Norde, W., "Proteins at Interfaces." Veenman and Ronen, Waginingen, 1976. 5. Norde, W., Furusawa, K., and Lyklema, J., in "Proceedings, Sixth International Congress on Surface Active Substances," p. 209. SpringerVerlag, Munich, 1972. 6. Lyklema, J., and Norde, W., Croat. Chem. Acta 45, 67 (1973). 7. Nyilas, E., in "Colloid and Interface Science" (M. Kerker, Ed.), Vol. 5. Academic Press, New York, 1976. 8. Kim, S. W., Lee, R. G., Adamson, C., and Lyman, D. J., Polym. Sci. Technol. 7, 69 (1975). 9. Kim, S. W., Lee, R. G., Adamson, C., and Lyman, D. J,, Amer. Chem. Soc., Div. Org. Coatings Plast. Chem. Pap. 33, 341 (1973). 10. Walton, A. G., and Hurwitz, F. M., Biophys. J. 25, 57a (1979). 11. Mathot, C., and Rothen, A., J. Colloid Interface Sci. 31, 51 (1969). Journal of Colloidand Interface Science, Vol. 72, No. 2, November1979
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WALTON AND MAENPA
12, Morrissey, B., and Stromberg, R., J. Colloid Interface Sci. 46, 152 (1974). 13, McMillin, C. R., and Walton, A. G., J. Colloid Interface Sci. 48, 346 (1974). 14. McMillin, C. R., Ph.D. Thesis, Case Western Reserve University, 1975. 15. McMillin, C. R., and Walton, A. G., Fed. Proc. Fed. Am. Soc. Exp. Biol. 33, 1473 (1974). 16. McMillin, C. R., Saito, H., Ratnoff, O. D., and Walton, A. G., J. Clin. Invest. 54, 1312 (1974). 17. Walton, A. G., Gershman, H., and Soderquist, M. E., Croat. Chem. Aeta 50, 197 (1977). 18. Maenpa, F. C., and Walton, A. G., Biophys. J. 21, 148a (1978). 19. Hasselbach, W., and Heinberg, K. W., J. Membrane Biol. 2, 341 (1970). 20. Watkins, R. W., and Robertson, C. R.,J. Biomed. Mater. Res. 11, 915 (1977). 21. Anderson, J. M., Gibbons, D. F., Martin, R. L., Hiltner, A., and Woods, R., J. Biomed. Mater. Res. Symp. 5, 197 (1974).
Journal of Colloidand Interface Science, Vol.72, No. 2, November1979
22. Van Kampen, C. C., Jones, R. D., and Gibbons, D. F., Biomat. Med. Devices Artif. Organs 6, 37 (1978). 23. Fair, B. D., Saito, H., Ratnoff, O. D., and Rippon, W. B., Proc. Soc. Exp. Biol. Med. 155, 199 (1977). 24. Melous, B., Moravek, L., and Kostka, V., Fed. Eur. Biochem. Soc. Lett. 58, 134 (1975). 25. Behrens, P. Q., Spiekerman, A. M., and Brown, J. R., Fed. Proc. Fed. Am. Soc. Exp. Biol. 34, 591 (1975). 26. Chou, P. Y., and Fasman, G. D., Biochemistry 13, 222 (1974). 27. Walton, A. G., in "Textbook of Protein Structure," Chap. 10. Elsevier, Amsterdam/New York, in press. 28. Schiller, P. W., and Schwyzer, R., in "Proceedings, 1lth European Peptide Symposium, 1971," p. 354, North Holland Pub. Co., Amsterdam, 1973.
INTRODUCTION
The interactions between proteins and surfaces are of great importance in nature, probably beginning in the prebiotic stages of chemical evolution, where rock surfaces may have played a role in polymerizing and organizing primitive proteins or proteinoids (1). In the most sophisticated biological structures present today, proteins or glycoproteins play an important role at interfaces including cells, joints, and arteries. Indeed, it is widely believed that protein adsorption at the blood vessel wall plays an integral role in control or activation of thrombogenesis (2). Questions concerning the behavior of proteins at "foreign" or nonbiological surfaces have been brought forth by the need to develop blood-compatible materials and prostheses (3) but are also important in various forms of chromatography, and in all cases where animal or vegetable matter 1 Current address: U. S. Biochemical Corp., Warrenville H t s . , Ohio.
come into contact with plastics or other surfaces. Following insertion of a polymer or other material into blood, proteins, particularly serum albumin and fibrinogen, are adsorbed fairly rapidly. Blood clotting can be initiated by the activation of one or more blood clotting factors (proteins) on interaction with the surface and/or by stimulated release of proteins and other entities from blood cells as they interact with the surface. In view of the central importance of protein behavior at interfaces, considerable effort has gone into characterizing the process (4). Approaches to the study of proteins at interfaces have considered two main facets of the problem--how much and what type of protein is adsorbed and what, if anything, happens to the structure of the protein in the adsorbed state. The first problem may be approached conveniently by studying adsorption isotherms of one or more proteins on a welldefined substrate. Experimentally one is faced with the familiar problem in surface
265
Journal of Colloidand Interface Science, Vol. 72, No. 2, November 1979
0021-9797/79/140265-14502.00/0 Copyright © 1979by AcademicPress, Inc. All rights of reproduction in any form reserved.
266
WALTON AND MAENPA
chemistry of either providing a large enough surface area that adsorbs measurable amounts of protein, or finding a sensitive method or tag (usually radioactive labeling) that allows study of protein uptake on relatively small surface areas (of, say, polymer film). In an example of the former method Norde (5), Lyklema (6), and others have measured the uptake of serum albumin on polystyrene latex spheres by calorimetry. They find that an essentially irreversible uptake occurs, first to approximately monolayer coverage, and then beyond. They were unable to reach a definitive conclusion concerning structural changes of adsorbed protein. However, Nyilas (7), using albumin and a disperse silica substrate and calorimetry, concluded that adsorption from solution caused denaturation of the protein in the interfacial region. Kim and co-workers (8, 9) have measured the uptake of albumin and fibrinogen both singly and in competition on polymer films by labeling the protein with iodine. They find that the protein adsorbs to a monolayer and beyond but, in mixtures, the relative amount of each protein adsorbed is a function of the time of measurement and the nature of the substrate. Although radiolabeling of proteins provides a convenient method of measurement, there seems to be some concern that such a procedure can in itself cause minor changes in protein conformation a n d " stickine s s" (10). Another method of detecting the extent of protein adsorption at a flat interface has been ellipsometry (11). By this technique the extent of protein attachment is measured by assessing the thickness of the refracting layer. The technique is only well suited to smooth surfaces and particularly reflective surfaces and provides, at best, very limited information concerning the structure of adsorbed protein. Infrared differential spectroscopy has been used by Morrissey and Stromberg (12) to examine the attachment and conformation of bovine serum albumin (BSA) on silica. Conformational changes are difJournal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
ficult to assess for globular proteins in aqueous solution, but they concluded that no significant conformational changes occurred, even for low adsorbed concentrations in this system. In an attempt to provide more direct evidence of the conformation of adsorbed proteins on surfaces, we have previously presented a surface circular dichroic (C.D.) method in which the structure of a monolayer, or less, of an adsorbed protein was readily assessable (13). It was found that fibrinogen underwent no detectable conformational change for adsorption onto quartz, that there were slight changes with bovine serum albumin (14), and that there were considerable changes for the clotting activation protein, factor XII (Hageman factor) on the same substrate (15, 16). Whereas the surface C.D. method provides direct information on surface-adsorbed protein structure, it cannot be used if the substrate absorbs UV light strongly, is rough, is optically active, or cannot be cast in a thin film on a transparent substrate. To circumvent some of these difficulties, we have been exploring the application of fluorescence spectroscopy (17, 18). The potential advantages of such a technique are that it is very sensitive and has been used for determining environmental and structural changes in proteins, including those in cell membranes (19). In addition, since fluorescence occurs for most proteins in the 340-mm region and virtually no potentially useful substrate fluoresces at this wavelength, interference from the substrate is minimized. If the substrate absorbs ultraviolet light in the region of protein stimulation (280 nm) or emission, this effect can be corrected for. The sole disadvantage seems to be that, unlike C.D. spectroscopy, the data cannot be interpreted in terms of detailed structural changes. There are basically two approaches available for examining the fluorescence of adsorbed proteins. The first, expounded here, uses a particulate substrate and meas-
STUDY OF PROTEINS AT INTERFACES ures the amount of fluorescence quenching, i.e., the decrease in fluorescence during adsorption. This method uses the intrinsic fluorescence of the proteins which involves the fluorescence of tryptophan and sometimes tyrosyl residues in the protein chain. The second method proposed recently (20), involves internal reflectance fluorescence spectroscopy (IRFS) and can be applied to thin films coated upon reflectance crystals. This method involves measurement of fluorescence increase in the interfacial area but generally requires an extrinsic fluorophore. In the cited work (20), the adsorption of fluorescein isothiocyanate (FITC)-labeled y-globulin on silicone rubber films was measured. Each technique has advantages and disadvantages. In general it would be preferable to use a technique which relies on intrinsic fluorescence of proteins on limited-area films of reasonable thickness (>10 /~m). Neither method is capable of this resolution and indeed the electronic principles involved are different. The following report outlines some principles and experimental data for the adsorption of bovine serum albumin on particulate synthetic copolypeptide substrates. These random copolypeptides are similar to petroleum-based polymers in many of their properties and typically are based on L-glutamic acid and L-lysine as the acidic or basic components and L-alanine, L-leucine, and glutamic acid or lysine derivatives as the hydrophobic component. The structure of some of these units in a representation of part of the polymer chain is shown below.
T
-NH-C- CONH-C-CONH-
I (~H2)2
I CH3 L- olony[ residue
-eorbobenzoxy L-lysyl residue
|COOCH2 O 7 -benzyI-L-glufomyl residue
-NH-C-CONH-C-CONH-
] CH3
I CH3
NH
I
CBZ L-leueyl residue
267
This type of substrate was used because of its potential viability as a biomaterial (21, 22). Since polypeptides are in some ways similar to native protein materials, this type of substrate may act as a model biological interface. However, from an experimental point of view, the polypeptide could present particular difficulties since we seek to examine the adsorption (and conformation) of proteins consisting of a polypeptide chain, on a similar substrate. There has been a previous report from our laboratory of the adsorption of Hageman factor onto polypeptide substrates (23) in which it was shown that protein fluorescence is indeed modified by the substrate. The nature of the modification is explored in this work. EXPERIMENTAL The bovine serum albumin used in this work was Miles primary standard monomer. Copolypeptides for substrates were provided by Drs. J. M. Anderson and T. Hayashi and have viscosity molecular weights of 50,000 or more. Stock suspensions of amino acid copolymers were prepared by first dissolving 1 to 2 mg of polymer in about 2 ml of chloroform. A few drops of the resulting solution were shaken with about 50 ml of distilled water. The chloroform was driven off by gentle heating in a hot water bath with the aid of moderate evacuation by an aspirator. Care was taken to keep the chloroform emulsified by continuous shaking during the process. Once all the chloroform was driven off, the process was repeated until all of the polymer solution was used. The resulting aqueous suspension was kept under an aspirator vacuum for 1 hr to pull off the last traces of chloroform. The resulting suspensions were filtered through a coarse glass filter to remove any large particles that may have been present. Particulate suspensions were examined in a Coulter counter and an electron microscope (Hitachi Hu 11A). The particles were Journal o f Colloid and Interface Science, Vol. 72, No. 2, November 1979
268
WALTON AND MAENPA
20~
10C 0 Z
1'o
b
20 n r 2 x 5000,mm 2
4O
30
% t: 1000 Q.
u~ SO0
0
o
1'o
2'o
30
40
n r 2 x 5000, mm 2
FIG. 1. Histograms of particle surface area for polypeptide suspensions of Glu(OBzl)Z°:Leu 8° calculated from electron micrographs. (a) Histogram of particle surface area as a function of size for
Glu(OBzl)2°Leu8°. (b) Histogram of fraction of total surface area of Glu:Leu 1:4 vs particle surface area. Although there are a large number of smaller particles, their contribution to the total surface area is limited. essentially spherical but heterodisperse in size. Typical histograms are shown in Fig. 1, which represents the relative n u m b e r of particles that have a given surface area (estimated assuming a spherical shape), and the relative contribution to total surface area b y particles of a single size. The histograms were prepared from micrographs of 16 random segments o f the prepared grids. The particles showed the same X-ray diffraction and infrared spectrum as the bulk material (a-helix) and therefore have substantially the same structure as the bulk solid. Relative fluorescence intensity measureJournal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
ments were made using an A m i n c o - B o w man spectrofluorimeter with an Ortec photon counter. Adsorption measurements were made by first measuring the fluorescence of the protein solution in 0.001 M phosphate buffer at p H 7.0 and 25°C. The fluorescence of the particles in protein (in buffer) was measured on a separate sample (to avoid fluorescence bleaching) and the fluorescence of the remaining protein was measured from a third, equivalent sample after centrifuging. Prior to the actual adsorption experiments, a determination was made on the most effective centrifugation rates and times by using suspension blanks. The effective-
STUDY
OF PROTEINS
ness of the centrifugation parameters on separating the particles was evaluated by measuring the loss of scattered light intensity (in the spectrophotofluorimeter) of the centrifuged samples compared with a distilled water blank. Typically, less than 1% of the original suspension light scattered, and ultraviolet light absorbance remained after 2 hr of centrifuging at 20,000-30,000g. In these circumstances and at 4°C, no detectable amount of protein was removed from solution. Particles, with their adsorbed protein, were removed from protein solution by using the centrifugation method as described. Beam Attenuation Correction
Since the particles of the suspension are insoluble polypeptide derivatives, they adsorb and scatter ultraviolet light in the region of fluorescence excitation (282 rim) and emission (345 nm). Consequently the relative fluorescence intensity of the protein in the absence and presence of suspension would be different, even if no adsorption or other solution modification were achieved. This effect, beam attenuation, can be accounted for by measuring the ultraviolet spectrum at the two wavelengths, and introducing a correction factor. If, for example, the absorbance and scattering at 282 nm are 10% of the incident light intensity for a certain suspension in a 1 cm cell, then, because of the 90° scattering geometry of the fluorescence system, the average protein molecule in the fluorescence beam receives 5% less light. Similarly, if the absorbance and scattering at 345 nm are 10% of the emitted light, the correction is 5% based on the geometry of the instrument. Another way of explaining this correction is that light passes through only one-half of the cell until it is absorbed, and one-half (at right angles) after emission. The relation between fluorescence intensity and ultraviolet absorbance is discussed separately; however, we find that the loss of fluorescence intensity with
AT INTERFACES
269
various suspensions used here lies between 1 and 5%. Nature of BSA Fluorescence and Structure
Albumin is the most abundant protein in serum. Two slightly different albumins are in common use: human serum albumin (HSA), which consists of 585 amino acid residues; and bovine serum albumin, consisting of 581 residues, many of which are homologous with HSA. The primary sequences of both have been established (24, 25). There are two classes of fluorescing proteins, class A proteins that contain no tryptophan and class B containing one or more tryptophan residues. This distinction is made because, of the three amino acid residues that fluoresce, phenylalanine, tyrosine, and tryptophan, the first two are quenched in the presence of tryptophan. Thus for class A proteins the detectable fluorescence is due to tyrosine residues (the quantum yield of phenylalanine is very low) and class B protein fluorescence originates in tryptophan residues. Human serum albumin contains only one tryptophan residue (residue 214) and appears to be unique in that both tryptophan and tyrosine fluorescence are detectable. On the other hand, bovine serum albumin has two tryptophan residues (residues 134 and 212) and is strictly a class B protein (see Table I). On denaturation (i.e., disruption of the secondary and tertiary structure by heating) the fluorescence maximum shifts and decreases. For HSA it moves from 339 nm to 350 nm with a drop in quantum yield from 7.4 to 5.2 and for BSA there is a shift from 342 nm to 350 nm and a concomitant drop in fluorescence from 15.2 to 7.4. Clearly BSA fluorescence is more sensitive to conformational change (denaturation) than HSA and it is BSA that is used in this work. The BSA molecule is estimated to be 116 x 27 x 27 A from Langmuir trough measurements. The detailed three-dimensional Journal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
270
WALTON AND MAENPA TABLE I Comparison of Amino Acid Sequence in the Region of Tryptophan in Bovine and Human Serum Albumin BSA Asp Glu P-he] Lys Ala Asp 125
i
t
1
~
, kys kys iPhe~
Gly
I
130 I
~
= 135
*
140
HSA Thr A~, ffh_ej,i~ Asp A~nLG!,:Glu Th~'Zh_e:Leu Ly~',Syj_Tx~_L_~u_TL~_Le~_TL~_G1LUe 205
BSA
210
212
215
220
-Gfu-~u" "~rg-~a~ Leu rL}~--~a-[~, Set ~Va7-ATa-AFg-L~ -fer-gin'], Lys [Phe- P-to -ky~I
I
' : HSA .GIu . .GIu. Arg . . Ala . I. Phe . . Lys . .Ala. .
L
I
I
I
, , , . A1a . . iVal . . Ala . . Arg . . Leu . .Ser. .Glnl . Arg . . iPhe . . Pro . . Lys . .
structure of the albumins is not known; however, circular dichroism spectra indicate that >50% of the structure is a-helix in each case. There is internal homology in the structure, indicating that it probably consists of three equivalent structural units. The most unusual feature of serum albumin is the unique sets of C y s - C y s groups. These occur at residue numbers 90, 91; 168,169; 245, 246; 278, 279; 360, 361; 437, 438; 476, 477; and 558, 559. It is possible to perform a C h o u - F a s m a n type of conformational analysis (26) which shows, as expected, long
.
a-helical regions. Typically the sequence contains a large number of runs of the type -P-N-N-N-P-N-N-P-, where P is a polar residue (Arg, Lys, Glu, Asp) and N is nonpolar. Such sequences are typical of a-helices in which the - P - residues lie in contact with the solvent (water) and the remaining residues form a hydrophobic interior. With these features in mind it is possible to postulate a crude structure in two and three dimensions. Figure 2 shows a model for human serum albumin which is essentially a set of a-helices of length 72-82 residues (110-120 ~i) linked
Hyd rophobic interior
- - ~- helices
o
cys-cys
120A
~80 residues long
pairs
:::X
Irregular region
30A
,,
'
FIG. 2. Schematic representation of serum albumin, based on conformational analysis and known geometric properties. Journal of Colloid and Interface Science,
Vol. 72, N o . 2, N o v e m b e r 1979
STUDY
OF PROTEINS
271
AT INTERFACES
0.8
0.6
G)
0.4
O e., .O
ol e~ ,~ 0.2
i
2
4
6
B.S.A.
8
10
(//g/ml)
FIG. 3. Relative fluorescence intensity and absorbance of BSA in /zg/ml range. The line C corresponds to measured values; A is corrected for the exponential term in Eq. [3]. by di-Cys bridges. The hydrophobic N and C termini are imagined to form a hydrophobic lock down one side of the cylinder and residues 246-278 and 438-476 form a base of irregular structure. On this basis one has 3 8 6 - a - h e l i x bends/585 = - 6 5 % ahelix. The structure has a hydrophobic " h o l e " down the middle, capable of carrying fats or other hydrophobic material (one of albumin's main functions). The Trp residues in H S A and BSA lie in regions that are probably o~-helical. The model will serve as a reference but should not be taken too literally until substantiated by further evidence. Fluorescence occurs when an electronic excited state cannot dissipate its energy through excited vibrational and/or rotational states, i.e., by thermal decay. Indole ring systems, of which tryptophan is an example, fall into this category. Absorptive transitions o c c u r between the ground state (A) and first or second excited state 1La or 1Lb. It is construed that decay occurs by intersystem crossing to the 3La triplet state with fluorescence occurring by decay of that state to a vibrationally excited state (27).
The fluorescence observed at a constant exciting-light intensity is equal to the light absorbed multiplied by a factor q5 which is dependent upon the quantum yield and an instrumental factor: Fx -
6(lo - I)x
,
[1]
Io where Io is the incident and I the radiated or transmitted intensity. These factors may be related by the B e e r - L a m b e r t equation
axLc = 2.303 loga0 (Io/I)x,
[2]
where eX is the molar extinction coefficient, L the path length, and c the concentration. Then Fx = ~b(1 - e x p [ - e a c L ] ) .
[3]
Thus for small absorbance the B e e r - L a m bert relation should hold with fluorescence replacing absorbance. Figure 3 shows that Eq. [1] holds for BSA up to a concentration of - 5 /zg/ml; the method is accurate to 0.1 /zg/ml or better, and thus provides a useful concentration assay. Journal of Colloid and Interface Science, Vol. 72, No, 2, November 1979
272
WALTON AND MAENPA 100
342 nm is quenched by the presence of (Glu(OBzl))8°Ala 2° copolypeptide particles. There is approximately a 25% reduction of fluorescence caused by the particles, but there is no indication of a wavelength shift to 350 nm that would be indicative of protein denaturation. The effect is similar to the previously noted decrease of Hageman factor fluorescence on polypeptide particles (23). Using the correction procedure given in the experimental section, it is possible to show that 1-5% of the reduction in fluorescence is caused by absorbance and/or by the polypeptide particle scattering. The remainder is due to a genuine loss of fluorescence of BSA protein in the presence of the glutamate copolymer. The decrease in fluorescence, AF, increases with increasing protein concentration and substrate particle density and is clearly identifiable with a surface interaction. In order to establish what fraction of the surface adsorbed protein was fluorescing, the particles were removed from solution by careful centrifugation, avoiding centrifugation of solution protein, as described in the experimental section. By this means, the loss in protein from solution could be ascribed to surface adsorption. Figure 5
8O
60 _= c
u.
~ if'
h 20
:l..'l
•'7 I
200
..... i
I
I
300
I
400
soo
~./nrn
F ~ . 4. F l u o r e s c e n c e of B S A ( ) in the a b s e n c e of (Glu(OBzl)8°:Ala2°); ( - - - ) in the p r e s e n c e of particles of the copolypeptide; ( . . . ) after r e m o v a l of copolypeptide particles with a d s o r b e d protein. RESULTS
Figure 4 shows the result of a typical run in which the fluorescence of BSA at
1.0
GL
1:4
.8
o
e~ <~'°
•
x
O
1= .4 xo
I--
,2 BSA
.4 in soln., mg/m I
.6
.8
1~0
FIG. 5. Typical adsorption i s o t h e r m s for B S A adsorbed on (Glu(OBzl)2°:Leu 8°) copolypeptide particles in the low coverage ( < 10%) m o n o l a y e r region b a s e d on fluorescence and electron m i c r o s c o p e data. (Symbols × a n d O refer to separate runs.) Journal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
273
STUDY OF PROTEINS AT INTERFACES T A B L E II
surface remodeling, aggregation, and perhaps surface poisoning by random entities. However, the most startling feature is that, if the surface-adsorbed protein is fluorescing, the total fluorescence should decrease after centrifugation. In fact it increases as indicated in Fig. 4. This discrepancy cannot be explained by particle absorbance of light and/or scattering, both of which have been measured carefully. For some reason (that will become apparent shortly) protein adsorbed on the surface and in the immediate area of benzyl-group-blocked polypeptide substrates, does not fluoresce. We have compared these experiments with particles of (Glu2°:Alas°) copolymer and with colloidal silica. In neither case is there a decrease in fluorescence even though protein is adsorbed. Consequently particles of copolymers Glu(OBzl):Leu, Glu(OBzl): Ala, and Lys(CBZ):Leu cause fluorescence quenching of BSA, whereas the Glu:Ala polypeptide and silica do not cause such a change.
Table of B S A A d s o r b e d on Copoly-(Glu[OBzl]8°:Ala 2°) Showing Strong and W e a k A d s o r p t i o n L a y e r s a
Total BSA /~g/rnl
Adsorbed BSA (calc) (/xg/ml)
23.75 95 237 713
9.4 23.0 46.8 113
Adsorbed BSA (cent)° (/zg/rnl)
BSA stripped~ (/zg/rnl)
7.0 16.2 20.5 54
2.4 6.8 26.3 59
a Equilibration with protein for 4 - 6 hr. b Cent. refers to material centrifuged with the particles. c On the a s s u m p t i o n that the stripped protein does not fluoresce at all in the p r e s e n c e of the polymer.
shows the low-coverage region in which the uptake of protein by surface is essentially a linear function of protein concentration. If the suspension is left to stand, the rate of protein uptake diminishes rapidly, dropping to about one-half in a period of 24 hr. We ascribe this feature to a combination of
0~ 300
0.~."0~0-
5mm cell
/
/ /
200
0
O
ofo ~
° ~ o ~
lOmm cell
100
/° I
I
lOOO
BSA
I
(/~glmg)
3000
FIG. 6. Relative fluorescence o f B S A in high-concentration region showing s e c o n d a r y scattering effects. Journal of Colloid and Interface Science, Vol. 72, No. 2, November 1979
274
WALTON AND MAENPA
1l
.13
.12
"'e
IJ.
I: O N ~
.11
.10
g .09,
i
~,
BSA
mg/ml
FIG. 7. Polarization ratio of BSA as a function of concentration. This information indicates that association of molecules does not occur at these concentrations. The absolute accuracy of the measurements is in the range 10-15% and no significanceis attributed to the scatter at low concentration. If the difference between the fluorescence of the protein in the presence of particles (corrected for light absorbance and scattering) and the fluorescence of protein after removal of the particles is attributed to a 100% quenching of protein influenced by the substrate, it is possible to calculate this amount of protein. Table II compares the amount of protein adsorbed with an additional amount not fluorescing in the presence of particles. Of course if the protein influenced by, but not adsorbed to, the particles is only partially fluorescing, then the quantity of material involved must be increased accordingly. DISCUSSION
Searching for an explanation of the fluorescence behavior of BSA in the presence of benzyl-esterified polypeptide particles, there seem to be three possibilities. The first is that the particles cause denaturation of adsorbed protein. H o w e v e r , such a postulate does not explain the increase in fluorescence after centrifugation. Journal o f Colloid and Interface Science, Vol. 72, No. 2, November 1979
Second, it seems possible that when protein molecules are crowded together on the surface or in the surface boundary layer, they undergo self-quenching. Such an effect could originate either in multiple scatter of light or in an energy-transfer process. There is some evidence that such a process can occur: At concentrations of BSA (>1 mg/ml) the fluorescence of BSA is attenuated. If the cell path length is halved, the fluorescence increases, indicating mutual interference effects (see Fig. 6). Consequently if high concentrations of protein occur in the interfacial region some attenuation and intermolecular quenching may occur. Crowding together of molecules could be a result of molecular association in the form of dimers, trimers, etc. Such an association can be tested by assessing the fluorescence depolarization. If molecules b e c o m e associated, their rotational diffusion is slowed and hence the rotation between absorbance and emission is changed. If molecules are associating there is an increased fluorescence polarization with increase in concentration. Figure 7 shows that there is only a small polarization of BSA
275
STUDY OF PROTEINS AT INTERFACES
solutions which is independent of the protein concentration. Similarly the introduction of particles does not produce a detectable increase in polarization of fluorescence. The fact that the quenching effect is not seen with colloidal silica seems to indicate that neither of the above effects is the major contributing process. The third possibility is that of molecular fluorescence quenching. If there is an energy acceptor within range, it is possible for the tryptophan residues to lose electronic energy by energy transfer. The efficiency of energy transfer is given by E -
1 1 + (R/Ro) 6
,
[51
where R is the separation of donor and acceptor and R0 is a characteristic distance. Energy transfer generally is detectable in the rangeR = 10-100 A. Literature reports have concentrated exclusively upon the transfer of energy within the same molecule, for example, between Trp and e-N-dansyllysine in adrenocorticotropin hormone. The dansyl group acts as an excellent fluorescence acceptor (28). In the present case we propose that energy-transfer processes can occur between the Trp residue of BSA and the benzyl radical of the substrate. To our knowledge energy transfer between a protein and solid substrate has not been documented previously. In addition to the energy transfer with acceptor groups in the substrate, it is conceivable that energy transfer can occur between protein molecules in close proximity to one another, either on a surface or in an adjacent layer. According to the F6rster theory of energy transfer, the separation distance R is related to the molecular parameters by R =
K2 8.79 × 10--28JAD n 4
1 -- T E ]1/6
× 4 ~ ' ~
.
[6]
~ LySCBZ2O 0Leu8 ~"~~.._ a,.oBzrOA,a~O , !
I
200
300
t 400
,~- (nrn)
FIo. 8. Ultraviolet spectra of Glu(OBzl)8°:Ala2° and Lys(CBZ)2°:Leu s° showing the split 1Lo transition in the 258-nm region and absorbance in the 30-nm+ region.
TE is the transfer efficiency given by
TE=
KT Kr + KQ + KR
K~,, KF, and KQ are the first-order rate constants for nonradiative transfer of excitation energy, fluorescence emission, and internal quenching of radiation, respectively. JAD is an "overlap integral" which accounts for the overlap between absorbance and fluorescence spectra of the acceptor. The quantity 4~°ois the fluorescence quantum yield of the donor without energy transfer and K / n ~ is a factor accounting for the relative orientation of donor and acceptor. Of the preceding factors it is readily possible and important to establish the overlap integral JAD. If the fluorescence spectrum of Trp and the absorbance spectrum of the Journal of Colloid and Interface Science, Vol. 72, N o . 2, N o v e m b e r 1979
276
WALTON AND MAENPA 80
Glu(OBzl):Glu does not associate with BSA. Since the average separation of molecules at the concentrations used is > 1000/~, there is no effective energy transfer. On the other hand, the polycation polylysine interacts with the polyanion B SA; the Lys(CBZ): Lys copolymer should also complex. Figure 9 shows the fluorescence spectrum of the basic copolymer with BSA; clearly the fluorescence is heavily quenched. A study of the conformation of the components BSA and poly-L-lysine by difference C.D. spectroscopy shows no net change of conformation on mixing (see Fig. 10). There is, therefore, no concurrent denaturation of the protein that could account for fluorescence quenching.
6O ta e-
=
-/ c
,~ 40
•~
2o
~,
/
I
250
I
350
I
450
M.m FIG. 9. Fluorescence spectra of water-soluble polypeptides (a) Glu(OBzl)6°:Glu4° and BSA, and (b) Lys(CBZ)5°:Lys5° and BSA (1:1 weight mixtures in each case). substrate do not overlap, then JAD ---- 0 and no energy transfer can occur. Figure 8 shows the absorbance spectrum of Glu(OBzl)S°Ala 2° and Lys(CBZ)2°:Leu s° in the 300-nm region. To benzene and benzyl groups are generally ascribed three major bands in the 185- to 195-nm (aBa), 205- to 225-nm (1La), and 255- to 275-nm (1Lb) regions.The 1Lb band of the two copolypeptides centered at 258 nm slightly overlaps the Trp excitation band in the 300-nm region. H o w e v e r , there is a forbidden transition ~Ba,b for benzene in the 360-nm region (not seen in the polypeptide spectra) that has almost complete overlap with the BSA fluorescence spectrum. In order to test whether energy transfer is indeed the predominant mechanism of energy transfer, we h a v e prepared modified samples of Glu(OBzl)~°°-x:Glu x and Lys(CBZ)l°°-ZLys x that are water soluble. In solution we look for fluorescence quenching caused by the association of the polypeptides with BSA. T h e acidic p o l y m e r Journal of ColloM and Interface Science, V o l . 7 2 , N o . 2, N o v e m b e r
1979
I.U ,.,I
o c&d
a E 0
o U,I a
200
210
220
230
WAVELENGTH
240
250
(nm}
FIG. 10. Circular dichroism spectra of (a) BSA (0.1 mg/ml); (b) poly-L-lysine (1 mg/ml); (c) addition of spectra (a) and (b); (d) spectrum produced by mixture of poly-L-lysine and BSA. Curve (a) has been reduced in scale for purposes of presentation. (The eUipticity of BSA at 220 nm is -18,000 deg.cm2/ dmole; polylysine at 220 nm is + 5000 deg. cm2/dmole.)
STUDY OF PROTEINS AT INTERFACES
277
We conclude, therefore, that the origin not scatter light unduly or absorb ultraof fluorescence quenching of BSA on poly- violet light in the 300- to 600-nm region is peptide particles lies predominantly in required. Suitable substrates include incharge transfer from the molecules adhering organics (glass, etc.) or organics low in to the surface, and those in the interfacial double bonds and devoid of benzyl or conregion (particularly within a distance of jugated benzene groups. In the case where 100 A). It seems likely that if the molecule the substrate does act as an energy-transfer lies flat on the surface the fluorescence will agent, some idea of the surface environbe heavily quenched since both tryptophan ment can be obtained by determining deresidues will be in close proximity to the tails of quenching processes. Polymers substrate acceptor. On the other hand, per- expected to act as energy-transfer systems pendicular adsorption is likely to cause a would include polystyrene, polyphenyl, etc. greater separation of Trp and substrate, with decreased energy transfer. ACKNOWLEDGMENTS The loose layer, which is swept free on We wish to acknowledge the experimental assistance centrifugation, could consist of perpendic- of Mr. Mark Soderquist and Dr. K. Raina. The ular molecules and others in a surrounding polypeptides were supplied by Dr. James Anderson "methoric" layerY In any case the Trp at Case Western Reserve University and Dr. T. moieties in these molecules must lie within Hayashi of Kyoto University. Modified polymers were 100 A of the surface for quenching to occur. produced by Dr. J. Caplin. This work was carried CONCLUSIONS
The fluorescence data point to the formation of a strongly adsorbed set of BSA molecules that probably are flat on the surface. Other molecules are less strongly adsorbed and form a "methoric" layer. The concentration of loosely attached molecules is at least as high as that of strongly attached molecules. The mechanism of energy transfer is probably via a combination of the triplet (3Ba,o) and 1Lb states of the substrate and interaction with adjacent bound or unbound molecules. No evidence has been found of denaturation of BSA on adsorption on these substrates (though this is not excluded). We conclude that fluorescence measurements can only determine directly whether conformation changes accompany adsorption when there is no mechanism for energy transfer with the substrate. A particulate suspension of substrate that does 2 The term "methoric layer" has been widely used in the colloid literature by the Yugoslav school to mean dense surrounding layers of counter'ions. In this case we use the term to indicate dense interfacial layers of protein.
out under the auspices of a Program Project in Biological Materials N 1HLB 15195. The junior author (F.C.M.) was supported by an N.I.H. Traineeship Award. REFERENCES
1. FOX, S. W., and Dose, K., "Molecular Evolution and the Origin of Life," p. 196. Freeman, San Francisco, 1972. 2. Salzman, E. W., Blood 38, 509 (1971). 3. Brash, J. L., and Lyman, D. J., in "Chemistry of Biosurfaces" (M. L. Hair, Ed.), p. 177. Dekker, New York, 1971. 4. Norde, W., "Proteins at Interfaces." Veenman and Ronen, Waginingen, 1976. 5. Norde, W., Furusawa, K., and Lyklema, J., in "Proceedings, Sixth International Congress on Surface Active Substances," p. 209. SpringerVerlag, Munich, 1972. 6. Lyklema, J., and Norde, W., Croat. Chem. Acta 45, 67 (1973). 7. Nyilas, E., in "Colloid and Interface Science" (M. Kerker, Ed.), Vol. 5. Academic Press, New York, 1976. 8. Kim, S. W., Lee, R. G., Adamson, C., and Lyman, D. J., Polym. Sci. Technol. 7, 69 (1975). 9. Kim, S. W., Lee, R. G., Adamson, C., and Lyman, D. J,, Amer. Chem. Soc., Div. Org. Coatings Plast. Chem. Pap. 33, 341 (1973). 10. Walton, A. G., and Hurwitz, F. M., Biophys. J. 25, 57a (1979). 11. Mathot, C., and Rothen, A., J. Colloid Interface Sci. 31, 51 (1969). Journal of Colloidand Interface Science, Vol. 72, No. 2, November1979
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12, Morrissey, B., and Stromberg, R., J. Colloid Interface Sci. 46, 152 (1974). 13, McMillin, C. R., and Walton, A. G., J. Colloid Interface Sci. 48, 346 (1974). 14. McMillin, C. R., Ph.D. Thesis, Case Western Reserve University, 1975. 15. McMillin, C. R., and Walton, A. G., Fed. Proc. Fed. Am. Soc. Exp. Biol. 33, 1473 (1974). 16. McMillin, C. R., Saito, H., Ratnoff, O. D., and Walton, A. G., J. Clin. Invest. 54, 1312 (1974). 17. Walton, A. G., Gershman, H., and Soderquist, M. E., Croat. Chem. Aeta 50, 197 (1977). 18. Maenpa, F. C., and Walton, A. G., Biophys. J. 21, 148a (1978). 19. Hasselbach, W., and Heinberg, K. W., J. Membrane Biol. 2, 341 (1970). 20. Watkins, R. W., and Robertson, C. R.,J. Biomed. Mater. Res. 11, 915 (1977). 21. Anderson, J. M., Gibbons, D. F., Martin, R. L., Hiltner, A., and Woods, R., J. Biomed. Mater. Res. Symp. 5, 197 (1974).
Journal of Colloidand Interface Science, Vol.72, No. 2, November1979
22. Van Kampen, C. C., Jones, R. D., and Gibbons, D. F., Biomat. Med. Devices Artif. Organs 6, 37 (1978). 23. Fair, B. D., Saito, H., Ratnoff, O. D., and Rippon, W. B., Proc. Soc. Exp. Biol. Med. 155, 199 (1977). 24. Melous, B., Moravek, L., and Kostka, V., Fed. Eur. Biochem. Soc. Lett. 58, 134 (1975). 25. Behrens, P. Q., Spiekerman, A. M., and Brown, J. R., Fed. Proc. Fed. Am. Soc. Exp. Biol. 34, 591 (1975). 26. Chou, P. Y., and Fasman, G. D., Biochemistry 13, 222 (1974). 27. Walton, A. G., in "Textbook of Protein Structure," Chap. 10. Elsevier, Amsterdam/New York, in press. 28. Schiller, P. W., and Schwyzer, R., in "Proceedings, 1lth European Peptide Symposium, 1971," p. 354, North Holland Pub. Co., Amsterdam, 1973.