Determination of reserve albumin-equivalent for ligand binding, probing two distinct binding functions of the protein

ANALYTICAL

BIOCHEMISTRY

Determination Probing

121,

395-408

(1982)

of Reserve Two Distinct

Albumin-Equivalent Binding Functions

ROLF BRODERSEN, SIGNE ANDERSEN, CHRISTIAN FINN EBBESEN,* WILLIAM J. CASHORE,? Institute Neonaiology.

for Ligand Binding, of the Protein

JACOBSEN, OVE SQNDERSKOV, AND SIGURD LARSEN*

of Medical Biochemistry, University of Aarhus, 8000 Aarhus C, Denmark; *Department Rigshospitalet. 2100 Copenhagen, Denmark: t Women and Infants Hospital of Rhode Brown University. Providence, Rhode Island 02908; and $Sigurd Larsen, Inc., Vestergade 84, 8000 Aarhus C. Denmark Received

September

of Island,

3, 1981

A method is reported for determination of albumin binding capacity for various ligands in 50-~1 sample volumes. A small amount of a radioactively labeled test ligand is added to the undiluted sample and the rate of dialysis of the free ligand into an identical sample without added ligand is measured. The reserve albumin-equivalent concentration is defined as the concentration of a standard albumin preparation which in buffered solution gives the same rate of dialysis and hence the same ratio of free/bound concentrations of the added ligand. It is shown that the reserve albumin-equivalent concentration, thus defined, is identical with the sum of concentrations of carrier species, each multiplied by the first stoichiometric binding constant of the test ligand to the carrier and divided by its first stoichiometric binding constant to the standard albumin. Determinations of this parameter are suitable for studies of the chemical potential and transfer affinities of individual ligands and for determination of interaction among several binding substances. Two test ligands have been used, monoacetyldiaminodiphenyl sulfone and diazepam. The former is bound competitively with bilirubin while diazepam engages another, independent binding function. The method can thus be used for separate determinations of the degree of saturation of two distinct binding functions of albumin. Complex mixtures of several carrier proteins with interacting ligands can be studied.

Serum albumin is capable of transporting a diversity of low-molecular-weight substances, such as fatty acids, drugs, bilirubin, tryptophan, and some inorganic ions. Variations of binding affinities and interactions of the ligands have been described (see review, (l)), especially in conditions of overloading of the albumin, such as uremia, treatment with multiple drugs, and prematurity. A complete description of binding equilibria in serum is therefore difficult, if not impossible at present. The recent finding of independent binding of two ligands to the albumin molecule has, however, opened new prospects. At least two distinct binding functions have been demonstrated, one dealing with bilirubin and several drugs, the second 395

binding most other drugs (2-4) while additional functions account for interaction with some exceptional drugs (5), with long-chain fatty acids, and with copper and calcium ions. The present paper presents a method for separate determinations of the availability of albumin with discrete binding capabilities, applied to two drug-binding functions, one of which is also responsible for bilirubin transport. The reserve albumin-equivalent concentration for binding of a specific ligand in an equilibrium mixture containing several carriers and bound substances is here defined as the concentration of a pure standard albumin which gives the same ratio of free/ bound concentrations of an added trace 0003-2697/82/060395-14.$02.00/O Copyright 0 1982 by Academic Press. Inc. All rights of reproduction in any form reserved.

396

BRODERSEN

ET AL.

amount of the specific ligand in a solution of identical temperature, pH, and salt concentrations. MATERIALS AND METHODS Analytical priociple. A radioactive test ligand is added to the sample. The concentration of the added ligand must be small in order to avoid a shift of binding equilibria of other ligands. The amount of free test ligand dialyzing through a membrane into another compartment is measured after a short period, 10 min. In order to avoid loss of any diffusible, competitive ligands, dialysis is performed against an identical sample without added ligand. Dilution of the sample is kept at a minimum, and temperature, pH, and salt concentrations are controlled. Equilibrium of dialysis is far from being established during the IO-min period (Fig. 1) and the amount of dialyzed radioactive substance depends upon the initial rate of dialysis. This rate is proportional to the concentration of free test ligand which again is a function of the concentration of available albumin. The ratio of radioactivities in the two compartments is calculated and is independent of quenching which is equal on both sides. A standard graph is prepared, using pure albumin solutions and plotting the ratio of count rates as a function of the albumin concentration. Measured ratios from samples are entered in the graph whereby reserve albumin-equivalent concentrations are obtained. It is a necessary condition that the concentration of the test ligand is small compared to that of the carrier and further that binding equilibrium with the test ligand is established within a short period of time, compared to the time of dialysis. These conditions are usually fulfilled with samples containing large amounts of albumin as the carrier. It would be theoretically possible to apply the experimental principle to samples containing other carriers, even to mixtures of unknown substances without albumin. We

, Oo

100

>

Tune Imlnutesl

FIG. 1. The amount of [‘%]MADDS present in the right dialysis compartment, relative to the total radioactivity, as a function of dialysis time: 0, without alumin; 0, with 300 pM albumin present in both compartments. Phosphate buffer, pH 7.4, 37°C. The increase of radioactivity in the right compartment closely follows a first-order course until about two-thirds of the process has been completed, i.e., for more than IO min in the experiment without albumin. A slight deviation from first order seen in the later part of the course is ascribed to the fact that the membrane is not infinitely thin compared to the chamber width and is further related to the shape of the compartments. The initial rate of the process is proportional to the concentrations of unbound MADDS, with or without albumin present.

may still talk of the reserve albumin-equivalent concentration for binding of a test ligand in a sample, meaning that albumin in this concentration could replace all carrier species in the sample and bind the test ligand equally tight. In such cases it might be difficult to ensure that the above conditions are fulfilled and errors could result so that varying values of the reserve albumin-equivalent concentration would be obtained with varying dialysis time. The present paper deals exclusively with species containing albumin in high concentrations as the dominant carrier and results are independent of varying dialysis time. Apparatus. The dialysis chambers consist of two halves of polyacrylamide, fixed on either side of the membrane (Fig. 2). The volume of each compartment is 20 ~1, not

RESERVE ALBUMIN

DETERMINATION

397

FIG. 2. Dialysis apparatus with compartment volumes of 20 ~1. One dialysis chamber, A, is shown in cross section in B. The thermostatic arrangement consists of an aluminum block, C-E, with six tunnels, D, to hold the chambers during dialysis. Pipetting is carried out while the chamber is placed on the apron, E, which is part of the thermostatic aluminum block and is heated from above by the bulbs, F. The temperature can be measured in various places by the electric thermometer, Cl.

including the volume of the filling channels. The membrane is cut, 2 X 4 cm, from cellophane dialysis tubing, Union Carbide Corporation, type 36/32, nominal thickness 0.0008 in. Membranes in the range of thickness 18.5-19.5 pm are selected, using a gauge from Cary, LeLocle, Switzerland, type TMU-50255/ 10215. A thermostatic aluminum block is used to keep the temperature constant, in case of clinical samples at 37.0 + 0.3”C. This has six tunnels, each accomodating one dialysis chamber. During the procedure of filling, one chamber at a time is taken from its tunnel and placed on the thermostatic apron where it is heated from above by two electric bulbs, OSRAMConcentra, Imp., 220-230 V, 100 W, placed about 30 cm above the apron. These bulbs are given half the nominal voltage and in this way a red light is obtained with a rel-

atively high yield of infrared irradiation and harmless to any bilirubin present in the samples. By vertical adjustment of the position of the bulbs the temperature in the dialysis cells at the time of filling and emptying can be regulated. The temperature at several points of the aluminum block and inside a dummy dialysis chamber is measured by an electric thermometer from Kamstrup-Metro A/S, Aabyhoj, Aarhus, Denmark, type 8721-244, with sensor Pt 100. Preparation of N-[4-[(I-aminophenyl)sulfinyllphenyl][ I -“Cl-acetamide, ([WIMADDS). ’ Thirteen milligrams (50 pmol)

4,4’-diaminodiphenyl sulfone (DDS) was dissolved in 250 ~1 fresh distilled dioxane. ’ Abbreviations used: MADDS, monoacetyl-4,4’diaminodiphenyl sulfone; DDS, 4.4’diaminodiphenyl sulfone; TLC, thin-layer chromatography.

398

BRODERSEN

Heating to about 50°C was necessary for dissolution. The solution was cooled to 20°C and added to an ampoule containing 250 PCi (6.3 pmol) [ l-‘4C]acetyl chloride and the ampoule was agitated vigorously on a Whirlimixer. The inside walls of the ampoule were rinsed with an additional 100 ~1 dioxane and agitation carried on for 5 min. A mixture of 25 ~1 concentrated ammonia solution (25%) and 25 ~1 water was added. All solvent was then removed by flushing with dry nitrogen above the reaction mixture. The product was redissolved in a mixture of 125 ~1 methanol and 125 ~1 chloroform. The solution was applied on three or four precoated TLC silica gel 60 plates (Merck, Darmstadt, W. Germany), 10 X 20 cm, layer thickness, 0.25 mm. The application was performed in 20 X 0.5-cm bands. The plates were developed with benzeneethyl acetate, 4:6 (v/v). Two bands could be observed under uv light, containing DDS and MADDS, in order of decreasing migration (6). The [14C]MADDS band (R, = 0.43, calculated relative to DDS) was scraped off and put on a column, 1 X 10 cm, with a sintered-glass filter and eluted with a mixture of chloroform-methanol, 1: 1 (v/v). The product was rerun on another plate using the same developing system. Finally, all solvent was evaporated and the product was redissolved in ethanol and dispensed in glass vials, each containing 8.75 nmol MADDS, 0.35 &i. The ethanol was evaporated and the vials sealed and kept in the refrigerator. The yields were 32-55% calculated on the basis of radioactivity, with a specific activity of 40 mCi/mmol. Chemical and radiochemical purities were tested in a chromatographic system of chloroform-methanol, 1: 1 (v/v). Radiochemical purity of the labeled reagent was further ascertained by comparing rates of dialysis with rates observed by spectrophotometric determination of nonlabeled MADDS (7). Identical rates were observed with dialyzed fractions of MADDS varying from 0.01 to 0.4. For use, the contents of one vial were dissolved in 7 ~1 ethanol; 70

ET AL.

11, 0.53 M, phosphate buffer, pH 7.4, was added and mixed. This radioactive MADDS solution was used within 4 days. “C-Labeled diazepam. Diazepam was obtained as a gift from Hofmann-LaRoche Ltd., Basel, Switzerland, and was purified by chromatography on precoated TLC silica gel plates developed with hexane-dioxaneacetic acid, 85:15:1 (v/v). Diazepam was eluted with methanol, dispensed in vials, 5.0 nmol (264 nCi) in each, dried, and sealed. For use, the contents of one vial were dissolved in 5 ~1 ethanol and mixed with 50 ~1 phosphate buffer, as above, and used within 4 days. Human albumin standard. Standard graphs obtained with MADDS and diazepam as test ligands are shown in Fig. 3. Human serum albumin from KABI AB, Stockholm, Sweden, was used as a standard for binding of MADDS, while the same albumin was used for studies with diazepam, in some experiments after defatting with charcoal in acid solution (8). A few other albumin preparations were tried and gave curves similar in shape but numerically different. The ratio of left-/right-side radioactivities, determined with human serum albumin from Sigma Chemical Company, St. Louis, Missouri, was 0.62 times that found with Kabi albumin, with MADDS as a test ligand, indicating that the Sigma preparation has a lower affinity, or a lower capacity, for binding of MADDS. Treatment of Sigma albumin with charcoal in acid solution improved the binding so that a factor of 0.83, relative to Kabi albumin, was obtained. Charcoal treatment of Kabi albumin results in defatting but did not significantly change the binding of MADDS. The choice of human serum albumin from Kabi as a standard is further supported by findings in sera from 37 healthy adults, male and female, showing lower reserve albuminequivalent concentrations for binding of MADDS, 0.6 to 0.9 times the concentrations of albumin, when measured relative to this standard (9). (Serum albumin concentra-

RESERVE

ALBUMIN

399

DETERMINATION

500

Standard

AlbumIn

Cone

, &-IM

FIG. 3. Standard graphs for determination of reserve albumin concentration for binding of MADDS, 0, and diazepatn. 0. The ratio of scintillation count rates in left/right compartments (ordinates) is shown as a function of standard albumin concentration (abscissa) in a buffered solution, pH 7.4, 37”C, dialysis time IO min. Human serum albumin from Kabi was used as a standard for work with MADDS, and the same preparation, defatted, was used for diazepam.

tions were determined by an immunological method (lo).) The standard albumin thus approximates albumin of adult sera with maximal MADDS binding. Bilirubin2 solutions. Bilirubin was obtained from Sigma Chemical Company, and was purified and crystallized from methanol-chloroform (11). This preparation contains almost exclusively bilirubin IX-(r( Z,Z). Four milligrams was dissolved in 0.2 ml 0.5 M NaOH, and 5 ml water and 75 ~1 1 M HCl were added. For work with diazepam, chloride-free solutions were used, prepared with 30 ~1 1 M phosphoric acid replacing HCl. Bilirubin solutions were mixed with albumin at pH 9 to avoid aggregation of bilirubin acid. The mixtures were subse’ Bilirubin is here used for unconjugated bifirubin cu(Z,Zj, the main species of bilirubin in the human onate and the toxic component.

IXne-

quently buffered to pH 7.4. Bilirubin solutions were handled under red light. Procedure. Determinations are usually done in triplicate. A 75-111 sample, or standard albumin solution, is mixed with 5 ~1 of a solution of the test ligand in concentrated phosphate buffer. Twenty-five microliters of the mixture is pipetted into the left compartment of the preheated dialysis chamber. One to one and a half minutes is allowed for equilibration of the membrane with the sample. Dialysis is then started by pipetting 25 ~1 of the same sample (mixed with buffer as above, but without added ligand) into the right compartment. After 10 min +-lo s the fluid in the left compartment is withdrawn, and the compartment is rinsed with 25 ~1 of the same buffer, diluted 1: 16. The residue of rinsing fluid is drained with a strip of filter paper and discarded. The fluid in the right compartment is then withdrawn; diluted

400

BRODERSEN

buffer is entered and left in the right compartment for 1 min to allow extraction of radioactive substance from the membrane and is finally withdrawn. The dialyzed samples are combined with the respective rinsing fluids in 2-ml plastic scintillation vials and 2 ml scintillation fluid, Ria Luma, from Luma Systems AG, Basel, Switzerland, is added. Counting is performed for 20 min in a Packard Tricarb liquid scintillation spectrometer. The dialysis chamber is rinsed again with diluted buffer, drained with filter paper strips, and reused, or is filled with diluted buffer, sealed with tape, and kept in the tunnel of the thermostat. Sources of error. Precision is optimal in the range of 50 to 400 pM reserve albumin equivalent, where the coefficient of variation is about 4% for a single determination when MADDS is the test ligand. For diazepam the optimal range is 50-600 PM reserve albumin equivalent with a 4% coefficient of variation. A 1“C temperature increase causes an error of -3% with MADDS and -6% with diazepam. Deviations of pH between sample and standard do not cause measurable errors with MADDS in the range of pH from 6.8 to 8.2 while a 10% increase of measured reverse for binding of diazepam is produced by an increase of pH of the sample over the same interval while pH of the standard is kept constant. Similarly, results with MADDS are insensitive to variations of salt concentrations whereas determinations of reserve for binding of diazepam increase by 20% when the phosphate buffer concentration is increased from 10 to 100 mM. A specific effect of chloride ions is seen on the binding of diazepam; an increase of sodium chloride concentration from 0 to 20 mM causes a 35% decrease of the measured reserve albumin-equivalent concentration. Long-chain fatty acids also influence the binding of diazepam considerably; 1 mol palmitate/mol albumin halves the measured reserve. Results for binding of MADDS are independent of chloride ion and fatty acid

ET AL.

concentrations within physiological concentration ranges. All the above figures refer to determinations with standard albumin solutions. Larger variations are encountered when competing ligands are present in binding equilibrium, sensitive to temperature, pH, etc. The two compartments of the dialysis chambers were tooled with a spherical cutter, 10 mm in diameter, to a depth of 1.20 +- 0.02 mm. With this tolerance, no variation of results from chamber to chamber could be demonstrated. Capillary creeping of the sample over the top of the dialysis chamber may cause sizable errors, but can be prevented by slight greasing of the surface. Results are not sensitive to incidental shaking of the chamber. Even rough handling during the process of filling and incubation produced no measurable errors. Increase of membrane thickness by 11% resulted in a relatively smaller decrease of dialysis rate, 4% only. This is probably explained by the avoidance of stirring. The fluid on both sides of the membrane remains stationary throughout the process of dialysis. Diffusion takes place as in a gel, and membrane thickness is therefore less critical than in determination of dialysis rate with stirring in both compartments. A slight dilution, by a factor 1.06, is introduced on addition of buffer with and without test ligand and causes a certain error. This is partly compensated for by diluting the standard albumin solutions similarly. Ethanolysislhydrolysis of MADDS was observed when this ligand was kept for several days in ethanol solution and resulted in very low results for reserve albumin equivalent since the ethyl acetate/acetate ion formed carries the radioactive label and remains unbound by albumin. Similarly, hydrolysis of MADDS in serum would cause errors due to nonbinding of acetate ions. Such errors would be discovered by decreasing results of triple determinations, due to

RESERVE

ALBUMIN

401

DETERMINATION

progressing hydrolysis after addition of the buffered MADDS solution to the serum. No tendency toward decreasing values has been observed in several hundreds of samples analyzed.

ond and following. We may then disregard all terms of second and higher order as well as the first-order term in the denominator and obtain Cbound(i)/Cfree

=

Cbound(St)/Cfree

Relation to carrier concentrations and affinities. The reserve albumin-equivalent

concentration for binding of a ligand, as defined in the introduction and as measured by the present method, is identical with the concentration of albumin with a vacant site for that ligand, provided the ligand is bound to one site with the same affinity as to the standard albumin, competing with other ligands in the sample, and provided allosteric effects are absent. This ideal condition is rarely, if ever, fulfilled since the ligand may be bound to more than one site on the albumin molecule or to other proteins, and the binding may be allosterically influenced by other substances bound to the same carrier molecules. In a general case, the thermodynamic meaning of the reserve albumin-equivalent concentration, as defined above, may be deduced as follows. The test ligand is bound reversibly to j carrier species. The ith species is present in the concentration Pi and several molecules of the test ligand may be bound to this with the stoichiometric binding constants K(,l, &, etc. The concentration of test ligand bound to this species is cb,,und(i)and the concentration of free test ligand is cfree.At equilibrium we have

pi

cfreeKi,I 1 +

CfreeKi.,

+

2CfreeKi,lKi,Z +

CfreeKi,lKi.2

+

’ ’ ’ +

* * *

[II

7

and similarly for a standard experiment the same concentration of test ligand,

THERMODYNAMIC RELATIONS OF THE RESERVE ALBUMIN-EQUIVALENT CONCENTRATION

Cbound(i) -=

PJ.1

’

It is now presumed that the test ligand is present in low concentration, so that cfreeKi,, is much less than unity, and further that the first binding constant is larger than the sec-

=

P x

Kst.~

3

with [21

when p is the reserve albumin-equivalent concentration, as defined above, and KS,,, is the first stoichiometric binding constant of the test ligand to the standard albumin. Since CboundW

--Ii

Cbound(i)

9

i=l

elimination gives

of cfree from Eqs. [l] P = ,$ f’t(Ki,JKst,~).

and [2] [31

The reserve albumin-equivalent concentration thus consists of a sum of concentrations of all carrier species, capable of binding the test ligand, each multiplied by the first stoichiometric binding constant of the test ligand to that species and divided by the first stoichiometric binding constant of the test ligand to the standard albumin. A carrier species in the sense of Eq. [3] may be albumin or any other macromolecule capable of binding the test ligand reversibly. If other ligands are present besides the test ligand, each stoichiometric species is reckoned separately, i.e., each combination of carrier and ligand molecules, irrespective of localization at sites. In serum, containing various fatty acids and other albumin-bound substances, the number of carrier species may be high. Relation to free ligand concentration and transfer affinity. The concentration of free

ligand in an equilibrium mixture can be determined by the present technique. A trace amount of the same substance, radiolabeled,

402

BRODERSEN

is added and the reserve albumin-equivalent concentration is measured by comparison with standard experiments, using pure albumin. The ratio of free/bound ligand concentration in the sample is then the same as in a standard experiment with the concentration, p, of albumin, 1 CfW CfreeGt) -=-= Cbound

Cbound(St)

p

x

KS,

’

when KS, is the first stoichiometric binding constant of the ligand to the standard albumin. The total concentration of the ligand in the sample is

ET AL.

equivalent concentration, px, for binding of the test ligand X, is measured in a solution containing a carrier, P, and a low-molecularweight substance, Y, different from the test ligand. One molecule of the substance Y can be bound reversibly to one molecule of the carrier with the binding constant Ky. The first, stoichiometric binding constant of the test ligand to the carrier is K,, to the complex PY is Kxcy,, and Kx,st to the standard albumin. The solution contains two species binding X, i.e., P and PY, and according to Eq. [3] we have Kx,Y, + [PY] F.

Px = [PI fi c =

Cfree

+

Cbound

X.3

,

from which we obtain cfree = c/(1 + P x Kst).

[41

The concentration of free ligand in a sample, containing other ligands and unknown carriers, can thus be obtained from its total concentration, if the reserve albumin-equivalent concentration is measured and the binding constant of the ligand to standard albumin is known. This may be utilized for determination of the transfer affinity of a protein-bound substance from one compartment to another, 1 +

PZ&

1 + p,Ks,

9

[51

if its concentrations in both compartments are known and the reserve albumin-equivalent concentrations are measured.

[61

xst

If the total concentration of the ligand Y is Y, and that of the carrier is P, we further have [P] = P - [PY], [71 and [Y] = Y - [PY], PI since the concentration of X is small. In the absence of Y, the reserve albuminequivalent concentration for binding of X to P is PX(O) = PWx/Kx.st). 193 When binding equilibrium of Y to P has been established, the ratio of occupied to total carrier is rY

= [ywp,

and the binding equation becomes

WI

Determination of energetic coupling of binding of two ligands. Reserve albumin-

rYP

Ky=[Pl[Yl=(P-ryP)(Y-ryP)~ 01

r

Combination

y = [(P + Y)K,

of Eqs. [6]-[9]

--$$-1=ry[2-1j.

A plot of experimental

+ l] - ([(P + Y)Ky 2PKy

+ 11’ - 4PYK:)“2

gives [ 111

values of the left

side of [ 1 l] against ry, the latter calculated from [lo], should thus give a straight line. KxCy,/Kx can be obtained from the slope of the line by adding 1. Deviation from linearity

RESERVE

ALBUMIN

may be expected at high concentrations of Y, due to binding of more than one molecule of this ligand to the carrier. If the binding constant of Y to the carrier is high, KY & 1 /(P + y>, it becomes unnecessary to know the value of KY since ry then approximately is Y/P. In conclusion, if the carrier concentration, P, is known and the reserve albumin-equivalent concentration for binding of X is measured in the absence of the other ligand, Kx/ Kx,sl can be found from Eq. [9]. Further, if the binding constant of Y is known or is high compared to 1 /(P + Y) and the reserve albumin-equivalent concentration for binding of X is measured with varying concentrations of Y, the fraction Kxcu,/Kx can be found from the slope of the curve at low concentrations of Y, using Eq. [ 111. This fraction is a measure of the mutual energetic coupling of binding of the two ligands, since, according to Weber (12) K

AG,,,=RTln~=RTln~.

KY.

K

[12]

KY

Use of a deputy test ligand. The present technique can, as shown above, be used for studies of certain aspects of binding and transfer of a ligand, and its interaction with binding of another substance, provided the ligand can be obtained in a radiolabeled form and is suitable for use as a test ligand. The theoretical basis for these uses is exact provided test ligand concentrations are small compared to the carrier concentrations and provided binding equilibrium is established within a time considerably shorter than the time of dialysis. A more general use of the technique can be made on certain approximative assumptions, studying binding of a substance, unsuitable as a test ligand, if a practicable deputy ligand can be found and applied as a radiolabeled test ligand. Bilirubin is bound so tightly to albumin that dialysis can hardly be demonstrated; it is only slightly soluble at the pH of blood

403

DETERMINATION

plasma and cannot be used as a test ligand. MADDS has previously been proposed as a deputy ligand for studies of bilirubin binding in plasma (7). The condition for using a deputy ligand must be that reserve albuminequivalent concentrations for binding of the deputy ligand approximate those for binding of the ligand to be studied. According to Eq. [3], this means that the first stoichiometric binding constant of the deputy ligand to every carrier species in the sample, relative to its binding constant to the standard albumin should approximate that of the ligand to be studied. This condition is probably fulfilled to a satisfactory level in case of MADDS as a deputy ligand for bilirubin (7) (see further Discussion). RESULTS Interaction of MADDS with Albumin

and Diazepam

Determinations of reserve albumin equivalent for binding of MADDS in a buffered solution of albumin with varying concentrations of diazepam are pictured in Fig. 4A, and reserve albumin equivalent for binding of diazepam with varying concentrations of MADDS in Fig. 4B. These data can be used for determination of the energetic coupling of binding of MADDS and diazepam, according to Eq. [ 1 l] and [ 121. Both of these ligands have binding constants, KY, close to 1 X lo5 M-’ (3,7), which is considerably more than l/(P + Y). The approximation, ry = Y/P, can thus be used, especially since the reserve albumin-equivalent concentrations in these cases seem to be independent of the concentrations of the second ligand for ratios Y/P well below unity; a small error on ry is thus immaterial. The slope of the regression line for points at diazepam concentrations less than 200 PM in Fig. 4A is 0.0015 f 0.023. Hence, Kxcyj/Kx = 0.9985 + 0.023, and AGx,u = -1 f 14 cal/mol. A similar result is obtained from the observed influence of the concentration of MADDS

404

BRODERSEN

ET AL.

Interaction of Bilirubin and MADDS Albumin

01 0

100

200 Diazepom

MADDS

300 Cone

Cone

I * LOO

,,uM

.,uM

FIG. 4A. Reserve albumin concentration for binding of MADDS as a function of varying concentration of diazepam in a solution of human serum albumin (Kabi), 300 PM, in phosphate buffer, pH 7.4, 37°C. FIG. 4B. Reserve albumin concentration for binding of diazepam as a function of varying concentration of MADDS in a solution of human serum albumin (Kabi), 100 pM; other conditions as in A. Nondefatted albumin and a standard curve for this protein was used to obtain uniformity of conditions in A and B.

on reserve albumin-equivalent concentration for binding of diazepam, as pictured in Fig. 4B, AGx,v = 0 + 25 cal/mol, in good agreement with the above figure. Typical values of the energetic coupling of binding of two ligands to one protein are of the order of +lOOO cal/mol ( 12). The present findings thus indicate that MADDS and diazepam are bound independently to human serum albumin.

with

Figure 5 shows the results of titrating human serum albumin with bilirubin. Reserve albumin-equivalent concentrations, p, for binding of MADDS, are pictured as a function of the concentration of added bilirubin, Y. At zero bilirubin concentration, the reserve albumin-equivalent concentration is equal to the albumin concentration, 600 PM, since the standard albumin preparation was used. On addition of increasing amounts of bilirubin, the reserve albumin-equivalent concentration for binding of MADDS decreases, approximately according to a straight line, p + Y = 600 PM. The slope is -1. In Eq. [lo] and [ 1 I], P is 6 X lop4 M and KY is of the order of 10’ M-’ and thus much larger than 1/(P + Y). Further, Kx = Kx,st and hence KxCy,/Kx = 0, indicating that MADDS is not to a measurable degree bound to a complex of albumin with one molecule of bilirubin. This result may also be expressed by saying that binding of one molecule of MADDS and binding of one molecule of bilirubin engage the same binding function of human albumin. In this simple case, a buffered solution of pure standard albumin and bilirubin, the reserve albumin-equivalent concentration for binding of MADDS is equal to the concentration of albumin without bound bilirubin, as seen from Eq. [3] which reduces to P=

[PI-

Interaction of Bilirubin and MADDS Infant Serum

with

Pooled serum from healthy infants’ cord blood was titrated with bilirubin, similarly as above. The result is seen in Fig. 6. Without added bilirubin, the endogenous bilirubin concentration was 21 PM and reserve albumin-equivalent concentration for binding of MADDS was 200 PM, which is low compared with the albumin concentration, 440 PM. Addition of increasing amounts of

RESERVE

ALBUMIN

405

DETERMINATION

8 0

::

\

:

8 0

L P .-c E 2 < a, : : c?

\ i \ 8

I

0 0

FIG. 5. Reserve albumin concentrations 600 PM, as a function of added bilirubin

I Bilirubin

I cont.,

for binding concentration,

bilirubin causes decreasing values of reserve albumin-equivalent concentration, initially following a straight line. By extrapolation, this line intercepts the axis of bilirubin concentration at 440 PM, which is the albumin concentration of the sample. Equation [9] gives Kx/Kx,s, = 0.48. In this case, KY, the binding constant of bilirubin to albumin of infant plasma, is not known but is presumably very large compared to l/(P + Y). A plot of (px/px(o) - 1) versus ry again has the slope -1 for small concentrations of bilirubin and we obtain Kxcy, = 0. The albumin of this pooled infant serum sample thus has a decreased binding affinity for MADDS but still binds MADDS and bilirubin competitively by the same function. In a number of infant sera, partly obtained from premature and sick patients, reserve albumin equivalent for binding of MADDS was measured by the present technique. For comparison, titrations with bilirubin were carried out, and monitored by the peroxidase method after dilution 1:40 ( 13). Results are seen in Fig. 7. Titration of serum with bil-

I ,uM

500

w

of MADDS in a buffered pH 7.4, 37°C.

solution

of albumin,

irubin for determination of the amount of available albumin after dilution 1:40, using the peroxidase method for indication of the end point, gave results in excess of the reserve albumin-equivalent concentration for binding of MADDS in the majority of samples, in some cases by a factor of 10. Interaction of Bilirubin with Albumin

and Diazepam

Figure 8 shows reserve albumin concentrations for binding of diazepam as a func-

FIG. 6. Reserve albumin concentration for binding of MADDS in infant serum, as a function of the concentration of added bilirubin. Albumin concentration in this sample was 440 pM, and that of bilirubin was 21 pM, pH 7.4, 37°C.

406

BRODERSEN

500,uM

MADDS

Method

FIG. 7. Comparison of bilirubin binding capacities (reserve albumin concentration), determined in 75 adult and infant serum samples, by titration with bilirubin, monitored by the peroxidase method after dilution I:40 (13) (ordinates), and measured in the undiluted sera, without added bilirubin, with MADDS as a deputy ligand (abscissa). Most of the observed points are located above the line, x = y, indicating that bilirubin titration after dilution of the sample gives higher values than the MADDS method, applied to undiluted serum.

tion of added bilirubin in a solution of standard human albumin. The slope is -0.003 f 0.019, indicating that bilirubin and diazepam bind independently, with an energetic coupling as low as 2 +- 12 cal/mol. In a buffered solution of standard albumin and bilirubin, the reserve albumin-equivalent concentration for binding of diazepam is thus equal to the albumin concentration and does not vary with varying bilirubin concentrations. This follows from Ki,l = Ks,, and Eq. [3] which gives

ET AL.

is bound solely to one site on the albumin and that allosteric effects from other ligands are absent. In the general case, when these conditions are not fulfilled, the reserve albumin-equivalent concentration indicates the concentration of a standard albumin preparation which in pure solution binds the test ligand equally tight as in the sample. This means that the results have a well-defined thermodynamic meaning, even when dealing with complicated samples, containing unknown carriers and mixtures of interacting ligands. Although fictitious concentrations are measured in such solutions, which may not contain albumin at all, the results are suitable for determinations of free ligand concentrations, affinities of transfer from one compartment to another, and energetic coupling of binding of two ligands, on an exact basis, as shown above. Studies using two test ligands, MADDS and diazepam, indicate that these substances are bound independently to human serum albumin while bilirubin engages the same binding function as MADDS and is bound independently of diazepam. The latter finding confirms previous results (3). The ob-

p = [P] + [PB] + [PB,] + . . - = P. DISCUSSION

A method has been presented for assessment of binding of a ligand in samples containing albumin or other macromolecular carriers and additional ligands in binding equilibrium. The results are given as reserve albumin-equivalent concentrations for binding of the test ligand and are identical with

concentrations of albumin with a vacant site for this substance, provided the test ligand

F G E E 03

I

I Bilirubin

I

I 500

0

Cone

'

,,uM

FIG. 8. Reserve albumin concentration for binding of diazepam as a function of varying concentration of bilirubin in a solution of human serum albumin, defatted, 600 pM, in phosphate buffer, pH 7.4, 37°C.

RESERVE

ALBUMIN

servations are consistent with the idea that albumin has two main binding functions, described by Sudlow et al. (2) as Sites I and II, and by Sjijhlm et al. (5) as the warfarin and diazepam binding sites. The majority of albumin-bound drugs can be classified as binding primarily to one or the other of these sites (5). On the other hand, conformational changes of the albumin molecule are important in the binding processes, so that a description of the albumin molecule as having a number of discrete, preformed sites or binding regions may not be entirely satisfactory (14,15), except for the copper ion (16). The term bindingfunction has accordingly been used in the present paper to allow for the possibility that ligand-induced conformational changes may be different according to type of ligand and may be so profound that a site for one ligand may not be recognized in the structure of the complex with another substance. Binding phenomena are here treated in a purely stoichiometric sense in order to render the conclusions independent of the possible presence or absence of preformed sites in the albumin molecule. Even with these reservations it is clear that albumin has at least two distinct binding functions, one dealing with bilirubin, warfarin, MADDS, and many drugs, and the second binding diazepam and most other protein-bound drugs. It would be tempting to hypothesize that the degree of occupation of albumin in serum from patients receiving multiple-drug treatment could be described for practical purposes by the concentration of reserve albumin equivalent, defined and measured as in the present paper, using one test ligand for each of the two main binding functions, e.g., MADDS and diazepam. The technique would appear suitable for further studies of this type. Clinical interest in the present method has focused on binding of bilirubin. This pigment in itself is not suitable as a test ligand, and MADDS has accordingly been proposed as

DETERMINATION

407

a deputy on the presumption that reserve albumin concentrations for binding of MADDS would be identical with those for binding of bilirubin (7). This requires that allosteric effects of other ligands on the binding affinity of bilirubin must be identical with those on binding of MADDS. This may well be the rule although notable exceptions have been found with laurate ( 17), and some tightly bound drugs, i.e., furosemide and ethacrynic acid, at high concentrations (7). These exceptions are without practical implication, since the allosteric effecters mentioned are found in serum at low concentrations. The basis for using MADDS as a deputy ligand for studies of the binding of bilirubin seems reasonably good, as judged from present evidence, although one should be prepared for certain deviations in the case of individual drugs. Clinical experience has shown that reserve albumin-equivalent concentrations for binding of MADDS are low in small and critically ill premature infants and are increased by exchange transfusion (18) by giving albumin ( 19,20), and by phototherapy (21, 22). The utility of the method for studying bilirubin-displacing effects of drugs in vitro as well as in vivo has also been demonstrated (9). Results of clinical determinations of the reserve albumin-equivalent concentration for binding of MADDS in newborns have been compared with those obtained by titration of available albumin with bilirubin, using the peroxidase method for monitoring the end point. Discrepancies are very considerable, as seen in Fig. 7, and may be explained by the dilution, 1:40, introduced in the peroxidase method, and by the use of stoichiometric amounts of bilirubin to titrate the available albumin, both of which would tend to diminish the effect of competitive ligands in the samples, thus explaining the high reserve albumin obtained by titration with bilirubin. This potential source of error should be weighted against the possible sources of

408

BRODERSEN

error in using MADDS as a deputy ligand for bilirubin. Calculations of the affinity of transfer of bilirubin from blood plasma to the brain, as it occurs in kernicterus, are possible on the basis of reserve albumin-equivalent determinations for MADDS (and thus for bilirubin), if bilirubin is deposited as the insoluble acid in the brain tissue (23,24). This possibility is being clinically evaluated as a means for assessment of one risk factor in neonatal hyperbilirubinemia (18-22,25). Other risk factors may be related to changes of the blood brain barrier, determining the kinetics of transfer. Another method for correct determination of available albumin, utilizing a spin-labeled selective trace ligand, has been introduced by Hsia et al. (26). ACKNOWLEDGMENTS We wish to thank Inger Bonnevie, Karen Cameron, Grethe Brodersen, and Anne Marie Bundsgaard for technical work, and Frede Nielsen for the drawings. J. Jacobsen, Ph.D., has kindly reviewed the manuscript. The following grant support is gratefully acknowledged: Danish Medical Research Council 5 I2- 10767, 5 l210626, and 512-15538 (R.B. and F.E.); Nordisk Gjenforsikrings Selskabs Jubilaeumsfond, (F.E.); Diabetes Research Center Grant HD 11343-01, National Institute of Child Health and Human Development, NIH, U. S. A. (W.J.C.).

ET AL. Brodersen, R., Sjijdin, T., and Sjoholm, 1. (I 977) J. Biol.

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in proof: Considerable variation of membrane permeability has been observed, especially when the cut membranes were kept for several months, exposed to daylight. Errors can be avoided by calibration, relative to a daily standard. Note added

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