©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Lipopolysaccharide Binding Protein-mediated Complexation of Lipopolysaccharide with Soluble CD14 (*)

Peter S. Tobias (§) , Katrin Soldau , Julie A. Gegner , Douglas Mintz , Richard J. Ulevitch

From the (1) Department of Immunology, The Scripps Research Institute, La Jolla, California 92037

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES

ABSTRACT

Endotoxin (lipopolysaccharide; LPS) activates a wide variety of host defense mechanisms. In mammals LPS binding protein (LBP) and CD14 interact with LPS to mediate cellular activation. Using sucrose density gradients and a fluorescent endotoxin derivative we have investigated the mechanism of LPS binding to LBP and the soluble form of CD14 (sCD14). LPS binds to LBP to form two types of complex; at low ratios of LPS to LBP complexes with one molecule of LBP and 1-2 molecules of LPS predominate, while at high ratios of LPS to LBP a large aggregate of LBP and LPS predominates. Complexes of LPS with sCD14 do not form large aggregates, consisting of only 1-2 LPS bound to a single sCD14 even at high multiples of LPS to sCD14. LBP catalyzes LPS binding to sCD14. Catalysis by LBP apparently occurs because LBP provides a pathway for LPS to bind to sCD14 which avoids the necessity for LPS monomers in aqueous solution. The dissociation constants for LPSLBP and LPSsCD14 complexes were determined to be 3.5 10 and 29 10 M, respectively. These numbers suggest that when LBP and sCD14 are present at roughly equal concentrations as they are in normal human plasma and compete for limited LPS, the LPS will predominantly associate with LBP.


INTRODUCTION

The exposure of organisms as diverse as horseshoe crabs and mammals to endotoxin (lipopolysaccharide, LPS)() results in the activation of a wide variety of host defense mechanisms. Recent studies of mammalian cells have described a unified pathway for the LPS-dependent activation of many cell types including monocytes, macrophages, neutrophils, endothelial cells, smooth muscle cells, and some epithelial cell lines. The common elements of the pathway include LPS binding protein (LBP), a 60-kDa plasma glycoprotein, and CD14, a 55-kDa glycoprotein. On monocytes, macrophages, and neutrophils, CD14 is present as a glycerophosphoinositol tailed membrane (mCD14) component (1) . In the presence of LBP, LPS binds to the mCD14 and initiates cellular activation (2, 3) . However, other LPS-responsive cells such as endothelial cells, smooth muscle cells, and some epithelial cell lines do not express mCD14. Instead, they have a receptor for complexes of LPS with the soluble form of CD14 (sCD14), which circulates in plasma without a glycerophosphoinositol tail (4, 5, 6) . Here again, LBP facilitates the formation of LPSsCD14 complexes.

In this report we describe the use of fluorescein-derivatized LPS (FITC-LPS) to permit continuous monitoring of the formation of LPSLBP and LPSsCD14 complexes. Heretofore it has not been possible to observe the formation of LPSLBP and LPSsCD14 complexes on a real time basis with a method that also permits the quantitative determination of kinetic and equilibrium parameters for formation of these complexes. We also describe our characterization of LPSLBP and LPSsCD14 complexes using sucrose density gradients. Others have used nondenaturing polyacrylamide gel electrophoresis to characterize LPS protein complexes (7) , but we found sucrose density gradients more useful since they permit more analysis of the separated components. Taken together, we conclude from these results that LBP catalyses the formation of LPSsCD14 complexes and suggest a mechanistic pathway by which this occurs.


EXPERIMENTAL PROCEDURES

Salmonella minnesota Re595 LPS was isolated and fluoresceinated with fluorescein isothiocyanate (Molecular Probes, Eugene, OR) by published methods (8, 9) . FITC-LPS from several commercial sources was not useful for these experiments. The LPS concentration of the derivative was determined by assay of the 2-keto-3-deoxyoctulosonic acid and measurement of the fluorescein content by optical density at 493 nm as described (9) . The products used here had substitution ratios of 20-30 mol % fluorescein. Based on the structure of S. minnesota Re595 LPS, its molecular weight was taken to be 2300 (10) . While the derivatization procedure used to make FITC-LPS may alter its physical properties from the unsubstituted molecule, the FITC-LPS and the [I]ASD-LPS retain their biological activity (data not shown). Rabbit LBP was isolated from acute-phase rabbit serum (11) . Human recombinant soluble sCD14 was isolated by immunoaffinity chromatography (Immunopure Protein G IgG Orientation Kit, Pierce, Rockford, IL) using monoclonal antibody 63D3 (American Type Culture Collection, Rockville, MD) from the supernatants of Chinese hamster ovary cells transfected, essentially as described in Ref. 12, with the cDNA for human CD14. To prepare S-labeled sCD14 and S-labeled sLBP CD14 or LBP, expressing Chinese hamster ovary cells were metabolically labeled with [S]Met (DuPont NEN, Boston, MA). The labeled proteins were prepared from 24-h culture supernatants by immunopurification using mAb 63D3 for sCD14 and mAb 18G4 for LBP. The mAb 18G4 as well as mAb 2B5, with specificity for human LBP, were kindly provided by D. Leturcq and A. Moriarty (R. W. Johnson Pharmaceutical Research Institute, La Jolla, CA). [I]ASD-LPS was prepared from S. minnesota Re595 LPS as described (11) . H-Labeled LPS and non-radioactive LPS from Salmonella typhimurium PR122(Rc) was obtained either from R. Munford (13) or commercially (List Biologicals, Campbell, CA).

Sucrose density gradients were centrifuged for 1 h 20 min at 4 °C in a pre-chilled TV-865 rotor (DuPont, Burbank, CA) at 55,000 rpm. The gradients were 4 ml of 5-20% sucrose in 0.05 M phosphate, 0.14 M NaCl, 2 mM EDTA, pH 7.4, layered onto 0.2 ml of 40% sucrose. The samples were incubated at 37 °C for 15 min and then chilled to 4 °C before 0.2 ml was layered on top of the gradient; aliquots of the samples were removed before centrifugation to determine total radioactive material for calculation of recovery. Typically, recovery of radioactive material from the gradients was 65-95%.

Fluorescence was monitored using an SLM 8000 fluorimeter (SLM Instruments, Urbana, IL) with excitation at 490 nm and emission at 520 nm. In plain glass cuvettes (7 mm 45 mm, Sienco, Morrison, CO) S-labeled LBP was observed to disappear from solution at a significant rate, presumably by adsorption to the glass (data not shown). In order to minimize binding of LBP and FITC-LPS to the cuvettes, they were filled with 10 mg/ml bovine serum albumin (Sigma) in water for 1 h after which they were extensively rinsed with water and stored moist at 4 °C. As indicated by the slow declines of fluorescence seen at long times in some figures, this procedure is not totally successful. Reactions were conducted in 50 mM phosphate, 100 mM NaCl, 2 mM EDTA at pH 7.4, except for the experiments reported in Fig. 3 b which were conducted in 10 mM phosphate, 2 mM EDTA, at pH 7.4 with KCl as required. Apparent pseudo-first order rate constants and dissociation constants were obtained from the fluorescence data using the curve fitting procedures of SigmaPlot (Jandel Scientific, San Rafael, CA). When loss of fluorescence due to adsorption to the cuvette was significant (see above), it was included as a time linear component of the overall fluorescent change. In some instances the amount of FITC-LPS approached 10% of the binding protein. Therefore, in all cases, Kvalues were obtained by fitting the data to a ``quadratic'' hyperbola rather than a ``rectangular'' hyperbola as defined by Equation 1,

On-line formulae not verified for accuracy

where F is the observed fluorescence, L and P are the concentrations of FITC-LPS and protein, respectively, SQRT is the square root operator, and F and K, the overall fluorescence change and the dissociation constant respectively, are the fitted parameters.


Figure 3: a, fluorescence intensity versus time of FITC-LPS (4.2 10 M) added at time A mixed with LBP (4.2 10 M) added at time B. Inset, the data from the larger figure and their fit to a pseudo-first order reaction with k = 4.72 10 s. b, the influence of salt concentration on the apparent first order rate constant for FITC-LPS-LBP formation. c, the net change in fluorescence upon mixing LBP with FITC-LPS (4.4 10 M). The solid line is fit to a quadratic hyperbola with K apparent of 1.0 10 M (see ``Experimental Procedures'' for details).



The photoaffinity probe [I] ASD-LPS (8 10 M) was incubated with several mixtures of sCD14 and LBP (see legend to Fig. 6) at 37 °C for 15 min before chilling to 4 °C for photolysis as described previously (11) . The proteins were then separated by SDS-polyacrylamide gel electrophoresis using a 10% acrylamide gel and detected by autoradiography after Coomassie staining to detect protein molecular weight markers (Sigma).


Figure 6: SDS-polyacrylamide gel electrophoresis gel analysis of [I]ASD-LPS labeled mixtures of sCD14 with LBP at 0.3 10 M using 8 10 M [I]ASD-LPS. The autoradiograph of the gel is shown together with the mobilities of purified LBP, sCD14, and the molecular weight markers bovine serum albumin ( 67) and ovalbumin ( 43).




RESULTS

Sucrose Density Gradient Analysis of LPSLBP Complexes

When H-labeled LPS or S-labeled LBP are separately subjected to sucrose density gradient sedimentation, their mobilities are quite different as shown in Fig. 1, panels a and b, respectively. The concentrations of H-labeled LPS and S-labeled LBP used, as well as the mole ratios of the incubation mixtures and the resulting sucrose density gradient peaks, are given in . As indicated by the mobilities of albumin and ovalbumin (data not shown), S-labeled LBP sediments with the mobility expected for a 60-kDa protein. However, H-labeled LPS has a mobility considerably higher than would be warranted by its molecular weight of 4000 (13) , indicating a high degree of aggregation. When H-labeled LPS and S-labeled LBP are mixed and incubated for 15 min at 37 °C before sedimentation, the mobility of the H-labeled LPS shifts to the mobility of LBP while the mobility of the S-labeled LBP does not increase (Fig. 1, panel c). This could be the result either of LPS binding to LBP or simply disaggregation of the H-labeled LPS. Since [I]ASD-LPS and LBP can be demonstrated to form a complex at lower concentrations than used here (see Fig. 6), we conclude that a complex of H-labeled LPS and S-labeled LBP has formed and that this complex has the mobility of S-labeled LBP. As the amount of H-labeled LPS used is increased, a high mobility complex H-labeled LPS and S-labeled LBP is formed ( panels c-e). However, the mobility of the complex is not determined by the absolute amount of H-labeled LPS used, but rather the H-LPS:S-LBP ratio, as is seen by comparing panel f with d and e. In summary, these data suggest that H-labeled LPS and S-labeled LBP may form two sorts of complex depending on the LPS:LBP ratio. One complex contains a single LBP and a small number (1, 2) of LPS molecules. The other contains a large number of LPS and probably multiple LBP.


Figure 1: Sucrose density gradients of mixtures of H-labeled [LPS and S-labeled] LBP. Sedimentation direction is from right to left. H-Labeled LPS and S-labeled LBP were mixed at the concentrations shown in Table I for 15 min at 37 °C before chilling to 4 °C for centrifugation. See text for methods.



Sucrose Density Gradient Analysis of LPSsCD14 Complexes

When H-labeled LPS, S-labeled sCD14, and LBP are mixed in various concentrations (see ) and subjected to sucrose density gradient sedimentation, an H-LPSS-sCD14 complex forms only in the presence of LBP, as shown in Fig. 2. As was the case with S-labeled LBP, H-labeled LPS and S-labeled sCD14 sediment distinctly differently in the sucrose gradients (Fig. 2, panels a and b). However, in contrast to the case with S-labeled LBP, no obvious H-LPSS-sCD14 complex is formed when the two are simply mixed (Fig. 2, panel c). Addition of a small amount of LBP does enable an H-LPSS-sCD14 complex to form (Fig. 2, panel d). With S-labeled sCD14, however, only one size of H-LPSS-sCD14 complex is seen despite the use of relatively very high concentrations of H-labeled LPS (Fig. 2, panels e and f) compared to the concentrations used to generate large complexes of H-labeled LPS with S-labeled LBP. The mole ratio of LPS:sCD14 in the complexes (see ) is distinctly less than that in the incubation mixtures when LPS is in excess. Both the size and composition of the LPSsCD14 complexes suggest that sCD14 can only complex with a limited number of LPS molecules.


Figure 2: Sucrose density gradients of mixtures of H-labeled LPS, S-labeled sCD14, and LBP. Sedimentation direction is from right to left. H-labeled LPS, S-labeled sCD14, and LBP were mixed at the concentrations shown in Table II for 15 min at 37 °C before chilling to 4 °C for centrifugation. See text for methods.



Enhanced Emission from FITC-LPS Upon Binding to LBP or sCD14

To develop a system that would permit real time determinations of the kinetics and equilibria governing interaction of LPS with LBP and sCD14 we asked whether the fluorescence signal of FITC-LPS would change upon complexation with LBP or sCD14. We observed an approximately 3-fold enhancement of fluorescence. The physical basis for this enhancement is not totally defined, but probably derives from relief of self-quenching in aggregates of FITC-LPS. S. minnesota Re595 LPS is undoubtedly strongly aggregated as are many types of LPS (14) . We observed that n-octyl--D-glucopyranoside also enhances the fluorescence, again suggesting that aggregate dissociation is responsible. As indicated by the sucrose density gradients of Figs. 1 and 2, incubation of H-labeled LPS with excess LBP or sCD14 reduces it's sedimentation velocity essentially to that of the protein, suggesting disaggregation of the H-labeled LPS.

Equilibria and Kinetics of FITC-LPS Binding to LBP

The time dependence of the fluorescence change when 4.2 10 M FITC-LPS reacts with a 10-fold excess of LBP (4.2 10 M) is shown in Fig. 3 a. The initial fluorescence is that of the buffer alone. FITC-LPS and LBP were added at points `` a'' and `` b'' respectively. The fluorescence change from 120 s, a time just beyond point b, to 250 s was fit as a first order process. The individual data points and the calculated first order curve are shown in the inset with k = 4.7 10 s. Experiments like these were conducted at a variety of FITC-LPS and LBP concentrations to characterize the kinetics of FITCLPSLBP complex formation. Over 2 orders of magnitude in [LBP], 4 10 to 4 10 M, k did not change significantly. These data suggest that formation of FITCLPSLBP complexes from LPS and LBP is a two-step process as shown in Fig. S1and that the molecularity of LBP does not change during the rate-limiting step. Because we can directly observe LBP bound to LPS aggregates (see Fig. 1) we favor a reaction scheme for FITC-LPS-LBP formation in which LBP rapidly binds to FITC-LPS aggregates. Subsequently, FITCLPSLBP complexes dissociate from these aggregates in the rate-limiting step.


Figure S1: Scheme 1



The rate of FITC-LPS-LBP formation was found to be strongly dependent on the salt concentration. The value of k was determined in 10 mM phosphate buffer at several concentrations of KCl as shown in Fig. 3 b. This observation is consistent with the idea that FITC-LPS-LBP formation involves transfer of hydrophobic structures such as the fatty acid tails of lipid A through a polar environment (15) .

The Kfor FITCLPSLBP complexes was obtained by analyzing the dependence on [LBP] of the difference in fluorescence between FITC-LPS alone and at the end of the association reaction. The data and their fit to an hyperbola are shown in Fig. 3 c from which K= 0.81(± 0.34) 10 M. The average of three determinations of Kwas 3.5 10 M with a range of 0.81 to 6.1 10 M.

As one would expect, unfluoresceinated Re595 LPS competes with FITC-LPS for binding to LBP (data not shown). Furthermore, monoclonal antibody 2B5, which binds LBP and blocks LPS binding to recombinant mCD14 expressing Chinese hamster ovary cells (16) also blocks FITC-LPS binding to LBP (data not shown).

Equilibria and Kinetics of FITC-LPS Binding to sCD14

Because complexes of LPS with either mCD14 or sCD14 are critical for activation of most LPS-sensitive cells, we investigated the reaction of FITC-LPS with sCD14. The data of Fig. 4 demonstrate the role of LBP in the binding of FITC-LPS to sCD14. At time A, 10 ng/ml FITC-LPS (4.5 10 M) was added to the cuvette, followed by 10 µg/ml sCD14 (1.8 10 M) at time B. The fluorescence does not change until time C, when LBP is added at 0.04 µg/ml (6.6 10 M). At the end of the ensuing fluorescence change, i.e. at about 300 s elapsed time, one may ask whether LBP has limited the reaction, since it is at the lowest concentration. Addition of another aliquot of LBP at C` does not cause a further fluorescence change, nor does addition of another aliquot of sCD14 at B`. Only addition of more FITC-LPS at A` causes a further increase in fluorescence. Since LBP was not limiting at the end of the first reaction, since almost no reaction occurred in its absence, and because 7 mol of FITC-LPS/mol of LBP bound to sCD14, we conclude that LBP must have catalyzed the binding of FITC-LPS to sCD14. In the experiments summarized in Fig. 5b, at the lowest LBP concentrations used nearly 500 mol of FITC-LPS bound to sCD14/mol of LBP.


Figure 4: Fluorescence intensity versus time upon sequential addition of 4.5 10 M FITC-LPS at A and A`, 1.8 10 M sCD14 at B and B`, and 6.7 10 M LBP at C and C`. Note that these concentrations are in the relative proportions of LBP(1):FITC-LPS(6.7):sCD14(268).




Figure 5: a, the net change in fluorescence upon mixing sCD14 with 4 10 M FITC-LPS in the presence of 5 10 M LBP. The solid line is fit to a quadratic hyperbola with K apparent = 2.9 10 M. See methods for details. b, the apparent first order rate constant for FITCLPSsCD14 complex formation versus LBP concentration with [sCD14] = 4.5 10 M and [FITC-LPS] = 8 10 M. The solid line is calculated for a rectangular hyperbola with k = 8.9 10 s and the [LBP] at which k = 0.5 k is 1.1 10 M. c, the apparent first order rate constant for FITCLPSsCD14 complex formation versus sCD14 concentration with [LBP] = 4.5 10 M and [FITC-LPS] = 8 10 M. The solid line is calculated for a rectangular hyperbola with k = 6.6 10 s and the [sCD14] at which k = 0.5 k is 2.5 10 M.



When the magnitude of the fluorescence changes such as those shown in the first part of Fig. 4 are measured as a function of sCD14 concentration, the Kfor FITCLPSsCD14 complexes can be determined to be 2.7 10 M as shown in Fig. 5 a. From four similar experiments, the Kdetermination averaged 2.9 10 M (± 30%). Changing the FITC-LPS concentration 4-fold, and changing the LBP concentration 2-fold had no observable effect on the value of the K. This is consistent with the sucrose density gradient data which suggest that LBP is not a constituent of the FITCLPSsCD14 complexes.

The kinetics of formation of FITCLPSsCD14 complexes as a function of LBP and sCD14 concentrations are shown in Fig. 5 , b and c. In the absence of LBP a rate constant of 1.7 10 s was observed. Fig. 5, b and c, suggest that as the protein concentration increases, there is a change in the rate-limiting step of the overall reaction, as discussed in the context of Fig. S1 .

Competitive Binding of LBP and sCD14 to LPS

As discussed above for Fig. 4, when LBP is at a low concentration relative to sCD14, the FITC-LPS transfers to sCD14. However, the determined values for the relative Kvalues of FITC-LPS-LBP and FITC-LPS-sCD14, 3.5 10 M and 29 10 M, respectively, suggest that as the ratio of LBP to sCD14 changes, the association of FITC-LPS with LBP and sCD14 should change. More specifically, at an LBP:sCD14 ratio of 1:8 the FITC-LPS should be approximately equally distributed between the two proteins. Since the fluorescence signals of the FITCLPSLBP and FITCLPSsCD14 complexes are essentially identical and will not permit verification of this prediction, we have used the radioiodinated, photoactivatable derivative of Re595 LPS, namely [I]ASD-LPS, to verify this prediction. Fig. 6 shows the relative labeling of LBP and sCD14 when 5 10 M (0.3 µg/ml) LBP and varying amounts of sCD14 are incubated and photolyzed with 8 10 M [I]ASD-LPS. As can be seen the labeling of the two proteins is roughly equal at an sCD14 concentration of between 2 10 and 5 10 M, verifiying the prediction made from the values of Kdetermined with FITC-LPS.


DISCUSSION

Heretofore it has not been possible to monitor the interaction of LPS with any LPS binding protein on a continuous, quantitative basis to explore the mechanism, kinetics, and equilibria of the reaction. While several strategems have been employed to determine binding affinities for LPS with several binding proteins (17, 18, 19, 20) , the kinetics of the binding reactions have not been measurable. Through the use of FITC-LPS, many, although not all, of the difficulties are removed. If, as we hypothesize, the increase in fluorescence derives from relief of fluorescein self-quenching in an FITC-LPS aggregate, then the method will not be useful for types of FITC-LPS which do not aggregate. However, these may be relatively few given the strongly amphipathic nature of the lipid A moiety of LPS. Additionally, not all types of LPS, and not all LPS partial structures, have readily derivatizable groups in appropriate parts of the molecule. Indeed, we were unable to observe any fluorescence change when the FITC derivative of Escherichia coli 0111:B4 LPS bound to LBP (data not shown). Wistrom-Aurell et al.() have observed that the fluorescein in the FITC derivative of E. coli 0111:B4 LPS is probably more distantly removed from the lipid a moiety than is the fluorescein in the FITC-LPS used here; thus self-quenching in that derivative is probably less pronounced. Nevertheless, the opportunity to make any real time, continuous measurement of even one LPS binding to LBP and sCD14 has considerable value. The reactions between FITC-LPS and other LPS binding proteins such as bactericidal/permeability increasing protein, cholesterol ester transport protein, and Limulus anti-lipopolysaccharide factor are currently under study and will be described elsewhere.

We conclude that LBP catalyzes the reaction of LPS with sCD14 for the following reasons. If LBP were a stoichiometric partner in the final complexes formed, this should be visible in the sucrose density gradients of Fig. 2, but it is not. LBP should be labeled by [I]ASD-LPS in the presence of any concentration of sCD14, but, as shown in Fig. 6, it is not. And if LBP is a component of the FITCLPSsCD14 complexes, the concentration of LBP should alter the apparent binding constant obtained when sCD14 is increased, but it does not. Thus three lines of evidence argue that LBP is not a constituent of the final complexes. Furthermore, Figs. 3 and 5 b show that LBP accelerates the reaction and that it delivers anywhere from 7 to nearly 500 molecules of FITC-LPS to sCD14. Thus LBP accelerates the reaction, LBP is not consumed in the reaction, and it does not perturb the final equilibrium. Thus we conclude that it is fair to claim that LBP acts as a catalyst in the reactions described.

Two possible reaction pathways which are potentially consistent with the kinetic data for the reaction of aggregated FITC-LPS (FITC-LPS) with LBP are summarized in in reactions 1-4 of Fig. S1. If the pathway involving reactions 1 and 2 is operative, then LBP reacts with FITC-LPS followed by dissociation of complexes of LBP with monomeric FITC-LPS. In this sequence, reaction 2 would be the rate-limiting step since reaction 2 involves no change in the molecularity of the reaction and the rate of LPSLBP complex formation was observed to be independent of the LBP concentration. If the alternative pathway involving reactions 3 and 4 is operative, monomers of FITC-LPS dissociate from the FITC-LPS and then react with LBP. In this sequence reaction 3 would be the rate-limiting step. Between these two pathways for LPS-LBP formation, we favor the upper pathway 1 and 2 primarily for the following reasons. The first is that complexes of LBP with LPS aggregates are directly observable in the sucrose density gradients. Complex formation via 3 and 4 does not involve any buildup of LBP bound to LPS aggregates since reaction 3 is rate-limiting. Second, if LBP reacted only with LPS monomers, because it has a higher affinity for LPS than does sCD14, LBP should inhibit the reaction of FITC-LPS with sCD14, not catalyze it. Undoubtedly there is some low equilibrium concentration of LPS monomers (21) which could react directly either with LBP or sCD14. Presumably it is this pathway which provides for the spontaneous reaction of FITC-LPS with sCD14 with a rate constant of 1.7 10 s.

Fig. 5 , b and c, suggests that as the concentration of either protein changes, the overall reaction undergoes a change in rate-limiting step. This is typically observed when one step of a multistep reaction is subject to catalysis (22) . For example, given the foregoing discussion, in Fig. 5 c we propose that the rate-limiting step is ``5'' at low [sCD14] and ``1'' at high [sCD14]. Under this interpretation, the value observed for k from the data of both Fig. 5, b and c, should be approximately 4.2 10 s, i.e. the rate constant for dissociation of an FITCLPSLBP complex from FITC-LPS-LBP. The observed values are 8.9 10 s and 6.6 10 s, respectively. Given the scatter in the data and the other vagaries in these reactions which we have experienced, we feel that theory and experiment are in acceptable agreement. Thus we conclude that the pathway for the formation of LPSsCD14 complexes catalyzed by LBP is that which is shown in boldface in Fig. S1.

Biologically, these results have several implications. As noted previously, recent work has suggested a catalytic role for LBP in assisting the presentation of LPS to sCD14 and that catalytic role is clearly supported by these results, as is the conclusion that sCD14 only binds a small number of LPS molecules (7) . While one might wish to extrapolate and suggest that LBP also catalyzes the binding of LPS to mCD14 (7) , i.e. CD14 on monocyte or neutrophil membranes, recent results from this laboratory show that LBP remains associated with mCD14 bearing cells when the cells are incubated with LPSLBP complexes (16) . Thus while LBP plays a catalytic role in the formation of LPSsCD14 complexes for binding to non-mCD14 bearing cells, LBP does not have a strictly catalytic role in the presentation of LPS to mCD14 bearing cells. Hailman et al.(7) have shown that LPSsCD14 complexes at 500 ng/ml LPS and 3 µg/ml sCD14 are agonists for neutrophils in the absence of LBP. However, Weingarten et al.(23) have shown that 0.01 ng/ml LPS was a direct agonist for neutrophils in whole blood. Thus the presence of LBP can apparently make 4 orders of magnitude difference in the ability of LPS to activate neutrophils. Similar effects are seen with monocytes (24) and this may be related to association of LBP with the cells.

It is also known that the in vitro activation of peripheral blood monocytes is some 1000-fold more sensitive to LPS than is the activation of endothelial cells and it has been proposed that the LPS-sCD14 pathway for activation of LPS-sCD14-sensitive cells exposed to blood is probably secondary to their activation by LPS elicited monocyte derived cytokines (25) . This hypothesis is supported by the measurements of Kfor FITC-LPS complexes of sCD14 and LBP; sCD14 binds FITC-LPS some 10-fold less tightly than LBP.

The fact that FITC-LPS is actually transferred to sCD14 in our experiments despite the fact that sCD14 binds FITC-LPS some 10-fold less tightly than LBP is a result of the fact that sCD14 was present at much higher concentration than LBP. However, in serum (or plasma), LBP and sCD14 are normally present at roughly equal concentrations. One would thus predict that in serum, LPS would predominantly associate with LBP. During an acute-phase response, LBP concentrations are considerably elevated, more so than sCD14. Thus after an acute-phase response, the balance of LPS bound to LBP and sCD14 would shift further to LBP, further diminishing the ability of LPS to stimulate cells via the LPS-sCD14 pathway. Thus although an important role for LBP catalysis of LPS binding to sCD14 is readily demonstrable in vitro, the role of this reaction in vivo remains to be defined. Perhaps it will be found at extravascular sites of sCD14 synthesis where the ratio of sCD14:LBP could be quite different from plasma (26) .

  
Table: Fig. 1 sucrose density gradients: LPS and LBP


  
Table: Fig. 2 sucrose density gradients: LPS, sCD14, and LBP



FOOTNOTES

*
This work was supported by National Institutes of Health Grants AI32021, HL23584, AI15136, and GM28485 and an American Heart Association fellowship (to J. A. G.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked `` advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

This is publication 9103-IMM from the Scripps Research Institute.

§
To whom correspondence should be addressed: IMM-12, Dept. of Immunology, The Scripps Research Institute, 10666 N. Torrey Pines Rd., La Jolla, CA 92037. Tel.: 619-554-8215; Fax: 619-554-3289; E-mail: tobias@scripps.edu.

The abbreviations used are: LPS, lipopolysaccharide; LBP, LPS binding protein; sCD14, soluble CD14; FITC, fluorescein isothiocyanate; ASD, 2-( p-azido-salicylamido)ethyl-1,3`-dithiopropionyl.

Wistrom-Aurell et al., manuscript submitted for publication.


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