Protein Diffusion in Cell Membranes: Some Biological Implications

INTERNATIONAL REVIEW OF CYTOLOGY. VOL. 87

Protein Diffusion in Cell Membranes: Some Biological Implications MICHAEL MCCLOSKEY AND MU-MINGPo0 Department of Physiology and Biophysics, Universiry of California, Irvine, California Introduction ........ Protein Diffu ....................... A. Experimental Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Biological Scope of Measurements........................ C. Temperature Dependency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Short- versus Long-Range Diffusion E. Diffusional Constraints . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Implications in Membrane Biology. ........................... A. Pattern Creation and Maintenance ........................ B. Reactions in Two Dimensions.. .......................... C. Biological Signaling at the Membrane D. Self-Assembly and Sorting .............................. IV. Closing Remarks ..... ......................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.

11.

19 20 20 21 22 24 25 34 34 45 59 66

I1 12

I. Introduction In the overall economy of a living cell, what difference does it make whether membrane proteins are capable of rapid diffusion or immutably immovable? How significant is the observation that a given protein has a translational diffusion coefficient of, say, and not lop9 cm*/second? Current efforts to capture and quantitate the motion of proteins in membranes are predicated on the assumption that this phenomenon has important biological implications. Our purpose here is to review some of the bases of that assumption, to speculate further on how passive diffusion of membrane proteins contributes directly to certain biological functions, and to consider how cells might deal with some of the potentially negative consequences of protein lateral motion. The intent is to provide an interpretive overview rather than a comprehensive summary of all published work in the field. Interested readers may find other recent reviews useful (Cherry, 1979; McConnell, 1979; Shinitzky and Henkart, 1979; Jacobson, 1980; Edidin, 1981; Jacobson and Wojcieszyn, 1981; Peters, 1981; Schlessinger and Elson, 1981; Webb et al., 1982; Axelrod, 1983). In 1968-1969 the seminal observation that hydrophobic regions of nerve and 19 Copyright 0 1984 by Academic Press. Inc. All rights of reproduction in any form reserved. ISBN 0-12-364487-9

20

MICHAEL MCCLOSKEY AND MU-MINGPO0

muscle membranes are “low-viscosity liquid-like’’ bilayers sparked a revolution in cell biology which has profoundly influenced present concepts of membrane structure (Hubbell and McConnell 1968, 1969). During the intervening 15 years there has been an explosion of empirical and theoretical research dealing with molecular motion and structural dynamics in both model and biological membranes. As a result, our knowledge of characteristic kinetic and thermodynamic properties of simplified lipid and lipid-protein systems has advanced rapidly. On the other hand, our ability to explain the workings of biological membranes in terms of those pure physical-chemical properties has not kept pace. Consider the following as illustrations of this point: (1) On the basis of a number of observations one must infer that bilayer fluidity or some closely related property is essential for normal life processes. This has been apparent for some time, and while qualitative rationale abounds an unequivocal molecular explanation does not exist. (2) Except for the erythrocyte we do not know what structural constraints govern the slower than theoretical diffusion rates of many integral membrane proteins. (3) Where there is striking long-range order in the lateral distributions of different lipid and protein species, we do not know what generates and maintains this topography. (4)On a smaller distance scale, say I0.1 pm, the homogeneity of the separate classes of lipids and proteins in cell membranes is largely unknown. ( 5 ) Although cholesterol is a ubiquitous and prominent constituent of animal cell plasma membranes, and though there exists an impressive catalogue of peculiar effects attributable to cholesterol in model systems, we do not know the primary function(s) of cholesterol in membranes. These questions provide an inkling of the true structural complexity of biomembranes. It is our belief, however, that persistent mechanistic dissection of the physical-chemical aspects of membrane-mediated events will eventually yield major clues about how biological membranes work.

11. Protein Diffusion: Known and Unknown A. EXPERIMENTAL TECHNIQUES

Since the initial finding in 1970 that surface histocompatibility antigens on mouse-human heterokaryons rapidly intermix after cell-cell fusion (Frye and Edidin, 19701, our understanding of protein mobility in cell membranes has taken a circuitous path. In 1973-1974 the first reported measurements of the rate of lateral diffusion led to the belief that membrane proteins are capable of “rapid” diffusion, with lateral diffusion coefficients in the range of 1 to 5 X IOW9 cm2/second (Edidin and Fambrough, 1973; Po0 and Cone, 1974; Liebman and Entine, 1974). Subsequent results using the method of fluorescence recovery after photobleaching (FRAP) have until recently yielded a 10 to lo3 times slower

PROTEIN DIFFUSION IN CELL MEMBRANES

21

lateral diffusion coefficient for membrane proteins. Moreover, in nearly all measurements with this method, a substantial fraction of the protein has appeared immobile for the duration of an experiment, usually ca. 30 minutes. Besides FRAP, a variety of other techniques has been employed to monitor protein lateral diffusion: intermixing of membrane components after chemically or virally induced cell-cell fusion (Frye and Edidin, 1970; Fowler and Branton, 1977; Schindler et al., 1980b); lateral spreading of locally applied labels (Edidin and Fambrough, 1973); redistribution of membrane proteins after electric field induced migration (Poo, 1981); passive accumulation of specific receptors at a ligand template (Michl et al., 1979); recovery of channel activity in locally inactivated zones (Poo, 1982). Most of the quantitative results, however, have come from FRAP. In this method, a fluorescent molecule like fluorescein or rhodamine is covalently attached to a probe, e.g., an antibody, lectin, or hormone, and the cell is treated with this specific fluorescent probe to label a particular membrane protein. Fluorescence on the cell surface is then irreversibly bleached in one or more regions by brief exposure to a high-intensity laser beam tuned to the excitation maximum of the fluorophore. The rate of recovery of fluorescence at the bleached region(s) is used to calculate a diffusion coefficient (Dvalue), assuming that recovery is due to lateral diffusion of proteins from the nearby unbleached regions of the membrane. The extent of recovery for proteins in biological membranes is, as a rule, less than 100%. Some recent experiments excluded, typical D values fall in the range of to lo-'* cm*/second, about lo3 to lo5 times slower than an average soluble protein in aqueous solution. On the other hand, isolated integral proteins diffuse at close to the theoretical limit (ca. lo-* cm*/second) and with nearly complete recovery when incorporated into artificial bilayers composed of fluid phase phospholipids. Fluorescent lipid analogs are generally rapidly mobile (ca. l o p 8 cm2/second)in both synthetic and biological membranes. Recovery is usually high for lipid diffusion in both systems. B. BIOLOGICAL SCOPEOF MEASUREMENTS With few exceptions (Stuhmer and Almers, 1982; Young and Poo, 1983) protein mobility has not been studied in intact tissues; isolated single cells of four principal classes have been the preferred objects of study. These include various cells from the blood and lymph, primary animal cell cultures, established cell lines, and vertebrate photoreceptors. Consequently, little is known about the diffusibility of membrane proteins in intact multicellular animal tissues. As will be discussed in a later section, there is now evidence that intercellular interaction can provide a passive localization mechanism for proteins in the animal cell plasma membrane. Metcalf et al. (1982) have published the only direct measurement of protein lateral diffusion in plant cell membranes of which we are aware.

22

MICHAEL MCCLOSKEY AND MU-MING PO0

Although the number of membrane proteins whose mobility has been examined continues to grow, there are some conspicuous gaps. As a class, peripheral proteins have not received adequate attention. Cytochrome c mobility in mitochondrial membranes has been directly assessed for the first time only in the last year, although its capacity to serve as a freely diffusing lateral shuttle of electrons has been discussed for some time (see Section III,B,2). Other peripheral proteins whose biological function(s) may depend upon lateral diffusion are plastocyanin, ferredoxin, and ferredoxin-NADP reductase in chloroplast thylakoids, phosphodiesterase and GTPase complex in rod outer segments, and perhaps the G-unit of hormone-activated adenylate cyclase in a host of cell types. Some cell surface proteoglycans, e.g., heparan sulfate (HS), are believed to be anchored to the membrane by a hydrophobic polypeptide (Kjelltn et al., 1981) as well as by specific interaction of the carbohydrate moieties with other membrane associated components, e.g., fibronectin; recent reports on the colocalization of heparan sulfate and ACh receptors in membranes of developing muscle (Anderson and Fambrough, 1982; Bayne et al., 1982) have heightened interest in the lateral mobility of surface bound HS, and it is likely that HS-ACh receptor coaggregates are assembled by a diffusion-dependent process. Changes in the type and amount of surface exposed proteoglycans accompany a variety of biological events such as mitosis (Kraemer and Tobey, 1972; Kojima and Koizumi, 1974), phagocytosis (Cappelletti et al., 1980), lymphocyte blastogenesis (Hart, 1982), and malignant transformation (Sampaio et al., 1977). Purified glycosaminoglycans can exert both inhibitory and stimulatory effects on cell division and growth (Chiarugi and Vannucchi, 1976; Takeuchi et al., 1976), and the aberrent adhesive properties of cancerous cells may arise in part from altered expression of cell surface proteoglycans. Aside from the potential significance of lateral diffusion in specific regulatory and recognition/ adhesion functions suggested for proteoglycans, from a purely structural standpoint some knowledge of the motional properties of glycocalyx constituents seems desirable, simply because the cell coat is such an intimate part of most animal cell plasma membranes (Rambourg, 1971; Luft, 1976). C. TEMPERATURE DEPENDENCY In view of the widespread interest in phase behavior of lipid-protein systems and the dramatic effects that lateral phase separations can have on protein diffusion in model bilayers (Wu et al., 1978; Smith et al., 1980; Vaz et al., 1981; Peters and Cherry, 1982), it is surprising that few systematic studies on the temperature dependence of protein diffusion in biomembranes have been done. In light of the current emphasis on cytoskeletal control of protein diffusibility and the lability of microtubules at low temperatures (Olmsted and Borisy, 1973), the common practice of holding freshly isolated cells on ice prior to FRAP measure-

PROTEIN DIFFUSION IN CELL MEMBRANES

23

ments is also somewhat suspect. In fact, cell deformability-and presumably the degree of polymerization and/or cross-linking of cytoskeletal polymers-can change appreciably between 37 and 4°C (Lichtman, 1970; Petersen et al., 1982). Because cytoplasmic levels of metabolites like ATP (Schindler et al., 1980a; Edidin and Wei, 1982), membrane potential (Edidin and Wei, 1982), or other conceivably temperature-sensitive factors may influence protein mobilities, the issue of temperature assumes even further significance. This seems especially pertinent for those cells removed from a native environment where temperature is usually closely regulated about values near 37”C, i.e., warm-blooded animals and incubators. In isolated membranes, several enzymes and transport proteins show breaks in their Arrhenius plots at characteristic temperatures beneath the host organism’s normal growth temperature (Kimelberg, 1977; Quinn, 1981; Ohyashiki et al., 1982; Brasitus, 1983; Rimle et al., 1983). As an isolated fact this could be explained by processes unrelated to thermotropic transitions in the bilayer. For example, a fairly temperature-dependent pK of an ionizable group in the active site or an altered K, due to a directly induced shift in an equilibrium between two conformational states of the protein might explain it. However, there are correlated changes in partitioning of small lipid-soluble molecules, tumbling of fluorescent probes, EPR and NMR order parameters, intensities of characteristic Raman and infrared bands, etc., which many of us take as evidence for the onset or completion of lipid phase separations. There is thus a widely shared belief that abrupt changes in the apparent activation energies for some enzymatic reactions and transport processes of membrane proteins are caused by thermally driven lipid phase changes; it is logical to hypothesize that part of the reason for “unexpectedly” low D values of cell membrane proteins is that surrounding lipids are partially solidified or otherwise segregated at the usual experimental (room) temperature. Available evidence on the temperature dependence of protein diffusion in cell membranes provides little support for this hypothesis. Using biotin depletion and fatty acid supplementation to enrich the membranes of chick myotubes with either high melting (elaidic) or low melting (oleic) fatty acids, Axelrod et al. (1978) found no more than a threefold change in the lateral diffusion coefficient of ACh receptors upon shifting the temperature from 12 to 3 1°C-for elaidate enriched, oleate enriched, or control cells. Similarly, over the interval 5 to 37”C, Hillman and Schlessinger (1982) found a modest threefold decrease in the lateral diffusion rate of ligated EGF receptors on a human epidermal cell line. Petit and Edidin ( 1974) witnessed a curious biphasic temperature dependence for intermixing of cell surface antigens on mouse-human heterokaryons which they attributed to lipid phase separations. As the temperature was lowered from 42 to 5°C mixing rates first dropped, then peaked at 15”C, then fell monotonically. It was suggested that reduction of the total diffusion space by channeling of pro-

24

MICHAEL MCCLOSKEY AND MU-MING PO0

teins within fluid paths of a solid-fluid mosaic enhanced the macroscopic diffusion rates and resulted in faster mixing at 15”C, but subsequent theoretical calculations have cast some doubt on this interpretation (Saxton, 1982). If one can extrapolate from this limited sample to other proteins and cells, how is he to rationalize the apparently small temperature dependency of protein diffusion with the above mentioned indications of thermotropic transitions? It is possible that extrinsic constraints are so great that lipid phase changes produce little extra effect. A more intriguing possibility which has found recent experimental support is that living organisms may suppress solid to fluid transitions observed in isolated membranes by promoting lipid fluidity through currently unknown processes (Cameron et al., 1983). D. SHORT-VERSUS LONG-RANGE DIFFUSION The possibility that biomembranes are laterally inhomogeneous over short distances accentuates one shortcoming of the FRAP method, namely, that it is only suited for measuring “long-range” (11 pm) lateral diffusion. The best one can do is obtain an average over all the short-range or “microscopic” D values which may exist. This limited spatial resolution is significant for a few reasons. First, it would be useful to be able to directly measure lateral diffusion of proteins and lipids in small cells like bacteria and sperm, and also in organelles such as the Golgi complex, endoplasmic reticulum, and thylakoid or mitochondrial membranes. FRAP is in general not applicable to these relatively small systems (for exceptions see Schindler et al., 1980b; and Hochman et al., 1983). Second, chemical reactions and other collision-dependent protein-protein interactions (e.g., self-assembly of multisubunit enzymes or channels) are probably more directly related to local than to long-range diffusion. Thus, if a 6receptor appears completely immobile using FRAP, one cannot say that it is incapable of reacting with cyclase by collisional encounters, because the local, or FRAP invisible, mobility may be significant. Of course, evidence of rotational immobility and/or visible aggregates is a separate matter. Third, heterogeneous microscopic D values would provide an indication that proteins are diffusing through locally inhomogeneous environments, and this might be a useful complement to biochemical studies aimed at characterizing short-range membrane structure. Short-range lateral diffusion of lipid analogs has been estimated with magnetic resonance techniques (Popp and Hyde, 1982, and references therein), triplet-triplet annihilation (Razi Naqvi et al., 1974), and pyrene excimer fluorescence (Galla and Sackmann, 1974); however, it is unlikely that any of these collision-dependentmethods will be extended to include proteins. On the basis of a recent study using spin labeled lipids, Laggner (1981) suggests that sar-

PROTEIN DIFFUSION IN CELL MEMBRANES

25

coplasmic reticulum membrane contains local diffusion barriers holding about 50 lipid molecules. Several techniques for measuring the rotational motion of membrane proteins are now available (see Cherry, 1979; Johnson and Garland, 1981), and these offer two distinct advantages over FRAP methods. First, the immediate environment of a protein is reflected more directly by the rate of its rotational diffusion than that of its translational diffusion. Second, rotational diffusion is more strongly dependent on molecular diameters than is translational diffusion (Saffman and Delbruck, 1975; Cherry, 1979; Hughes et a l . , 1982). This latter makes rotation rates a valuable probe of small-scale protein-protein association (e.g., dimerization, oligomerization) .

E. DIFFUSIONAL CONSTRAINTS I . Slow Mobility and the “Immobile Fraction”-What Do They Mean? Much current interest in the field centers on interpretation of the slow mobility and apparent immobility of proteins in cell membranes. As noted earlier, diffusion coefficients of proteins in cell membranes as measured by FRAP methods have in general been much smaller than those expected for proteins in a fluid lipid bilayer. Furthermore, in most photobleaching studies fluorescence recovery is incomplete, often ranging from 20 to 70% of prebleached levels. Partial recovery presumably results from immobility of the remainder of the labeled protein, or “immobile fraction. Consistent with this interpretation is the finding that roughly 100%recovery is obtained after a few successive bleaches of the same region. In model phospholipid bilayers in the fluid state, intrinsic proteins usually have a very large mobile fraction and they diffuse nearly as fast as fluorescent lipid analogues, certainly within a factor of 5 (see references in Section 11,E,6). This finding is more or less consistent with the theoretical predictions of Saffman and Delbruck (1975) for diffusion in bilayer membranes, namely, that lateral diffusion coefficients are weakly dependent on molecular diameters and inversely proportional (roughly) to the length of the diffusing molecule within and normal to the bilayer. Hence, for a protein with bilayer-spanning segments the predicted D value is about half that for a typical lipid molecule, which spans only half the membrane. Yet in membranes of intact cells the difference in diffusion rates of lipid and protein is often apparently one to three orders of magnitude, a result clearly at odds with Saffman-Delbruck theory. Two limiting assumptions of the original Saffman-Delbruck formulation are that intrinsic bilayer viscosity is much greater than that of the bathing aqueous phase and that diffusing molecules do not project beyond the hydrophobic bilayer core. Typical estimates of viscosity for the hydrocarbon region of fluid lipid ”

26

MICHAEL MCCLOSKEY AND MU-MING PO0

bilayers range from 1 to 10 poise (P), while that of water or physiological buffers is about 0.01 P. For synthetic lipid membranes the first assumption is reasonable and the second is therefore of little consequence; but just the opposite situation may exist in biomembranes. Hughes et al. (1982) have extended the Saffman-Delbruck theory to include lateral and rotational diffusion rates for all relative values of the bathing and bilayer viscosities. When applied to proteins and lipids for which both rotational and translational D values are available, the new calculations are more consistent with a high bathing viscosity (ca. 1 P) and a relatively low bilayer viscosity (ca. 0 . 2 P). One gathers that hydrophilic protein segments protruding into the aqueous phase may experience much more viscous drag than those within the bilayer, a notion to which we will return later in a discussion about the role of cell surface glycocalyces in restricting protein mobility. It is generally felt that constraints imposed by extramembranous macromolecules account for the slower than expected protein mobility, and the prevailing emphasis is on the role of cytoplasmic constraints. This and other possible mechanisms will be discussed in the following sections. It is worth noting, however, that even if we can pinpoint the cause(s) of slow diffusion, the existence of an immobile fraction also demands clarification. The puzzling fact is that any protein, whether it is a homogeneous or heterogeneous molecular population, always appears to contain an “immobile” subpopulation when examined with FRAP. Is there an anchoring mechanism (say cytoskeleton connection) that always works on a fraction of all the proteins in the membrane? Or, is there perhaps a wide spectrum of diffusion rates for any given protein, with the D value measured by FRAP representing an average rate of detectable diffusion, and the “immobile” fraction representing that subpopulation whose diffusion is too slow to measure? Whatever the explanation it will make a great difference in terms of our ideas about protein diffusion in cell membranes.

2. The Cytoskeleton There is clear evidence that cytoskeletal structures can somehow impede the motion and alter the distribution of integral proteins. Studies purporting to document the effect are far too numerous to cite here. Discussed below are results from three systems where FRAP has been used to demonstrate a direct correlation between cytoskeletal alterations and changes in lateral diffusion rates. a. The Erythrocyte. For the erythrocyte there is compelling evidence that lateral motion of band 3 and other intrinsic proteins is severely retarded by the spectrin-actin-ankyrin matrix which subtends the membrane. In normal mouse erythrocytes band 3 has a lateral D value of 4.5 X lo-” cm*/second at 24”C, while in erythrocytes from mice with a hereditary deficiency of cytoskeltal components the corresponding value is 55-fold greater (Sheetz et al., 1982). In human erythrocyte ghosts a reversible 50-fold increase in the lateral D value and

PROTEIN DIFFUSION IN CELL MEMBRANES

27

a jump in the mobile fraction (10 to 90%) of band 3 results from manipulations of temperature (21 to 37°C) and buffer concentration which are believed to destabilize the cytoskeleton and specifically disrupt band 3-ankyrin interactions (Golan and Veatch, 1980, 1981). Polyphosphate compounds like ATP and 2,3-diphosphoglyceric acid which partially depolymerize the erythrocyte cytoskeletal network cause a moderate (ca. 5 X ) increase in the lateral D value of band 3 (Schindler et al., 1980a; Sheetz et al., 1982). Based upon their work with the erythrocyte Koppel et al. (1981) worked out a detailed mathematical model of integral protein diffusion which treats the cytoskeleton as a matrix with labile cross-links whose rate of formation and breakage sets the observed lateral diffusion rate of intrinsic proteins. Using this conceptual framework they obtain a result consistent with the erythrocyte viscoelastic mechanical properties measured by Evans and Hochmuth (1978). Hypotheses on the nature of band 3-cytoskeleton interactions have come full cycle in the last decade. Original electron microscopic studies led to the conclusion that most of band 3 is directly bound to cytoskeletal polymers (Pinto da Silva and Nicolson, 1974; Elgsaeter e t a l . , 1976; Yu and Branton, 1976). Cherry et al. (1976) then observed that essentially complete elution of spectrin from the erythrocyte membrane did not alter the rotational diffusion rate of band 3, and concluded that no direct linkage could exist between band 3 and the cytoskeleton. In subsequent work the same group discovered that proteolytic cleavage of a cytoplasmic portion of band 3 significantly enhanced its rotational diffusion, and that elution of ankyrin with salt treatments also released constraints on the rotational motion of band 3 (Nigg and Cherry, 1980). They concluded that as much as 40% of band 3 is directly bound to ankyrin, a component of the erythrocyte cytoskeleton which links spectrin to the membrane (Bennett and Stenbuck, 1979; Luna et al., 1979; Yu and Goodman, 1979). Consistent with this notion, Golan and Veatch (1981) found that selective proteolytic cleavage of ankyrin totally releases diffusive constraints on band 3, and that addition of the released ankyrin fragment to untreated ghosts causes a modest rise in lateral mobility of band 3. Bennett and Stenbuck (1979) also showed that solubilized band 3 binds to purified ankyrin. Hence, current evidence does not favor the hypothesis that purely spatial barriers limit band 3 lateral motion, although as Koppel et al. (1981) point out, steric hindrance may well be the major impediment to long-range diffusion of the “mobile” fraction of band 3. b. Anchorage Modulation. ‘‘Anchorage modulation” refers to the inhibitory effect of Con A binding on the patching of several antibody cross-linked membrane proteins in various cell types (Edelman, 1976). The effect is global in that binding of Con A to local regions will inhibit patching over the entire cell surface. Because patching is a passive diffusion-mediated process is has been assumed that Con A inhibits patching by reducing receptor diffusion rates. Anchorage of these potentially patchable proteins is presumably controlled by

28

MICHAEL MCCLOSKEY AND MU-MING PO0

cytoskeletal constituents assembled into a hypothetical “surface modulating assembly” (Edelman, 1976). Recently it was shown directly that localized binding of Con A to mouse lymphocytes reduces the mobility of cell surface IgG (Henis and Elson, 1981). In the presence of Con A the D value and mobile fraction determined by FRAP are about sixfold less than in untreated cells. As with patching inhibition, this effect is global and occurs at a threshold value of about 10% coverage. Above 10% the response is coverage independent and beneath it no effect is observed. The strength of this experiment lies mainly in the finding that colchicine and cytochalasin B act synergistically to return both the D value and mobile fraction of surface IgG on Con A-treated lymphocytes to control values. Here it departs significantly from many previous experiments which had failed to document enhanced diffusion upon treatment with chemicals known to depolymerize microfilaments and microtubules. In accordance with prior studies these pharmacological agents did not raise D values or mobile fractions above control values on cells not treated with Con A. One may surmise from this work that cytoskeletal elements are somehow participating in Con A induced immobilization of surface IgG; nevertheless, the experiment reveals nothing whatsoever about why IgG diffusion is restricted to 5.3 X 10- l o cm2/second on untreated cells-and that seems to be a major question. c. Blebs and Ballooned Cells. A recent intriguing result is that lateral diffusion coefficients of several integral proteins are remarkably increased in plamalemma “blebs” on various cell types and in “bulbous” lymphocytes;both structures are induced to form by chemical treatment with formaldehyde/ mercaptans, phallacidin, or Con A (Tank et al., 1982; Wu et al., 1982). From cytochemical staining it appears that the plasma membrane in these structures is lifted off the underlying cytoskeleton; coincident with this structural alteration is a change in lateral D values of ACh, Con A and LDL receptors from less than 10- l o to about 3 X 10- cm2/second. Fluorescence recovery after photobleaching is also complete. Why is diffusion much faster in “blebs” than in untreated plasma membranes? Webb and associates favor the interpretation that loss of cytoskeletal connections to the plasmalemma releases diffusive constraints in “blebbed” membrane, but recognize that other factors could be involved. Until a rigorous biochemical analysis of lipid, protein, and carbohydrate constituents of “blebbed” membrane and plasmalemma fractions of bulbous cells is performed, other interpretations remain tenable. For example, externally disposed, highmolecular-weight components of the cell coat could be excluded from “blebbed” regions or lost from bulbous lymphocytes. In fact, Sugrue and Hay (1981) have demonstrated that corneal epithelial cells generate membranous blebs when stripped of their extracellular matrix, and that readdition of certain extracellular matrix constituents causes the blebs to retract. We have found (McCloskey et al., unpublished) that when rat basophilic leukemia or Xenopus cells are treated with

PROTEIN DIFFUSION IN CELL MEMBRANES

29

glycosaminoglycan degrading enzymes or trypsin, large protrusions, or “blebs,” sometimes appear on the cell surface. One might speculate that they form where the cell coat has been thinned the most by enzymatic attack. Interestingly, in these protrusions the FRAP-determined lateral diffusion coefficients of IgE Fc and Con A receptors are rapid and recovery is nearly complete. While the above-mentioned ‘‘blebs” are artificially induced structures, blebs do arise spontaneously on cells from many types of animals, both in culture and in vivo. These are transparent, hemispherical protrusions about 2-5 p,m in diameter which form within seconds and then slowly recede (Albrecht-Buehler, 1981). The bleb interior contains ribosomes and other small inclusions but lacks filamentous structure; it is thought to be much more fluid than the remaining cytoplasm. In culture, blebs are most prevalent as cells flatten out after division or after being plated; they form a dynamic ferris-wheel array on cells in some developing amphibian embryos. It would be of interest to measure protein diffusion rates in naturally occurring blebs to see if constraints are relaxed as in the induced “blebs.” If so, it would imply that proteins are capable of sudden, large-scale changes in mobility during their residence on the cell surface.

3 . Intrinsic Diffusion Barriers Although extramembranous constraints are undeniably important, the possibility that lateral motion of integral membrane components is also hindered by structural heterogeneity of the bilayer or by direct interactions with other integral components has not been rigorously excluded. In bilayers consisting of a binary mixture of cholesterol and dimyristoyl phosphatidylcholine the coexistence of solid and fluid regions can dramatically influence long-range diffusion of fluorescent lipids (Rubenstein et a l . , 1980; Owicki and McConnell, 1980). Measured D values for the solid-fluid mixture differ by more than an order of magnitude from theoretical microscopic D values in either phase. Klausner and Wolf (1980) find that certain fluorescent lipid analogs are largely immobile (mobile fraction 25%) in a solid-fluid mixture of di-C,,- and di-(=,,-lecithin while other analogs are almost 100% mobile, and they interpret this as due to selective partitioning of the different probes into either solid or fluid domains. Although biomembranes probably do not often contain the percentage of solid phase lipids present in either of these model systems, solid-fluid phase separation is only one mechanism whereby intrinsic lipid organization could influence protein mobilities. Several cases of fluid-fluid immiscibility involving phospholipids are now known (for references, see Berclaz and McConnell, 1981). Given the selective lipid dependence and preferential lipid-protein association constants exhibited by proteins like cytochrome oxidase (Marsh et a l . , 1982) it is possible that some proteins would partition unequally between two fluid lipid phases. In this case the protein would diffuse rapidly within both phases but conceivably more slowly across the phase boundary. If the protein enriched phase happened to be dis-

30

MICHAEL MCCLOSKEY AND MU-MING PO0

persed then the protein would be partially trapped, and its long-range diffusion somewhat impeded. Another potential intrinsic constraint in some systems is binding between the pendant oligosaccharide groups of integral proteins and glycolipids. The rotational motion of spin-labeled ganglioside head groups is restricted by interaction with glycophorin in reconstituted membranes (Sharom and Grant, 1978). Protein lateral diffusion might also be retarded if the interaction were multivalent for both protein and l i p i d d u e to multiple sites per molecule or to glycolipid clusters like those suggested by Sharom and Grant (1977, 1978). It is unlikely that the above constraints will fully explain the slow mobility or immobility, but they do merit consideration as potential contributions. Intrinsic diffusion limits of a different nature are mentioned in Section II,E,6. 4. Difluusion of Ligand-Free and Ligand-Bound Proteins

One serious reservation we have regarding FRAP-determined diffusion coefficients stems from a problem inherent to the experimental protocol. As generally practiced, photobleaching measures the diffusion not of native proteins but of extrinsically labeled species. Binding of specific ligands to cell surface receptors can have large effects on their mobility and lateral distribution. Examples of ligand-induced clustering or dispersal include the opioid peptide receptors (Hazum et a l . , 1980), thyrotropin receptors (Avivi et a l . , 1981), p-adrenoreceptors (Henis et a l . , 1982), chemotactic peptide receptor (Niedel et al., 1979), and receptors for hormones like EGF and insulin (Schlessinger et al., 1978). Oliver and Berlin (1982) discuss several cases of ligand-induced receptor migration in leukocytes and propose an “entrainment” of the ligand-receptor complexes in actively propagated membrane ‘‘waves. ” There is also the distinct possibility that lectin receptors may globally modulate their own mobility by signaling cytoplasmic attachments upon binding (Henis and Elson, 198I ) . Aside from these specific and sometimes biologically important effects, we are concerned with a possible nonspecific effect due to ligand binding. In particular, does the attachment of a large probe like an antibody Fab fragment impede diffusion of integral proteins due to its interaction with the thick, carbohydraterich cell coat (glycocalyx) which appears to surround all animal cells? Several experimental results bear directly on this issue. First we note that in most FRAP studies fluorescently labeled antibodies, antibody fragments, lectins, or other macromolecular ligands have been used to label the membrane proteins. As previously stated, the D values and percentage recovery are consistently lower than for integral proteins in reconstituted systems or for rhodopsin in the disk membrane. When a small (MW 441) fluorescent antagonist was used to label padrenoreceptors in Chang human liver cells the mobile fraction had a high D value of 1.4 X lop9 cm*/second at 23°C (Henis et al., 1982). Diffusion coefficients of free, or nonliganded proteins can be determined by back diffusion after

PROTEIN DIFFUSION IN CELL MEMBRANES

31

in situ electromigration (Po0 er al., 1979) and postinactivation recovery of functional activity (Poo, 1982). Using these methods the diffusion coefficients for Con A and acetylcholine (ACh) receptors in embryonic Xenopus muscle cells were estimated to be in the range of 1-4 X cm2/second. Using postelectric field relaxation D. W. Tank et al. (personal communication) have estimated that unliganded LDL receptors on an internalization defective human fibroblast cell line diffuse at a rate of 1.1 X cm2/second (22°C). Using FRAP, this same group finds a D value of 1.4 X 10- cm2/second for the LDL-receptor complex at 21°C (Barak and Webb, 1982). They found no evidence for cytoskeletal disruption by the electric fields, and suggested that LDL particles may interact with extracellular matrix components and thus impede lateral motion of ligated LDL receptors. There is also more indirect evidence that ligand-free proteins may diffuse quite rapidly. For example, when murine macrophages are plated on an immunoglobulin G (1gG)-coated cover slip the IgG Fc receptors disappear from noncontacting portions of the plasma membrane. The process is not blocked by metabolic poisons but is prevented by soluble IgG. Assuming it is due to passive diffusion of receptors to the IgG “trap,” a high D value of about 2.5 X l o p 9 cm2/second was calculated for the free receptor (Michl et al., 1979, 1983). Although the effect of ligand binding on protein diffusion remains a debatable issue, its final resolution will have important implications not only in the interpretation of experimental results, but also for a number of biological processes, e.g., hormonal and immune responses.

5 . The Glycocalyx and Extracellular Influences In lipid-protein model systems, an artificial cell coat can have remarkable effects on both the binding and diffusion of macromolecular “ligands.” The affinity of wheat germ agglutinin (WGA) for lipid vesicles containing the integral protein glycophorin is increased tremendously by the presence of a mere monolayer of high-molecular-weight dextran (Ketis et al., 1980); WGA binding also appears to become cooperative when dextran is present. These effects are not unique to dextran: other high polymers and bovine serum albumin work just as well. Ketis and Grant (1982) also find that by butressing the relatively sparse cell coat of extensively washed human erythrocytes with a layer of adsorbed BSA they can substantially increase the number of high affinity WGA binding sites. Restriction of oligosaccharide rotational and translational mobility may be involved in this phenomenon. Indeed, spin labels on the terminal sugars of liposome-bound glycophorin report a 20% reduction in rotational diffusion rate when the liposomes are coated with dextran and immunoglobulin or BSA (Lee and Grant, 1979). When specific anti-nitroxide antibodies are bound to bilayers containing phospholipids with a nitroxide head group, the antibody-hapten complex diffuses at

32

MICHAEL MCCLOSKEY AND MU-MING PO0

the same rate the lipid hapten does (Smith et al., 1979b). This is in the absence of an added “cell coat.” When a monomolecular layer of BSA is added to a similar system the lateral diffusion rate of the antibody-hapten complex is an order of magnitude smaller than the lipid hapten itself (Hafeman et al., 1981). One cannot help but conclude that the more firmly attached layer of proteoglycans, the covalently bound oligosaccharides of integral proteins, the carbohydrate chains of glycosphingolipids and accumulated detritus at the surface of real cells would provide a more effective diffusion barrier than a monolayer of BSA. We have tested this possibility by using FRAP to quantitate mobility of plasma membrane proteins in living cells after partially removing the cell coat. Subsequent to mild treatment with hyaluronidase, chondroitin lyase ABC, and Flavobacterium heparinase, there is as much as a 20% increase in the mobile fraction of soybean agglutinin bound to cultured Xenopus muscle cells, although the D value is apparently unchanged (Liu et al., unpublished observation). Whether more complete removal of the same glycosaminoglycans or other glycocalyx constituents would further increase the mobile fraction remains to be determined. Rapid diffusion of integral proteins in spectrin-depleted erythrocytes appears inconsistent with the glycocalyx being a significant diffusional constraint; however, of cells which have been stained for cell coat material the erythrocyte seems to have the thinnest and least dense coat of all (Luft, 1976). Furthermore, directly bound, small-molecular-weightfluorophores rather than antibodies have usually been used to label the erythrocyte proteins for diffusion studies. For further discussion of the glycocalyx, see Section 111,A,6. 6 . What Determines the “Viscous Limit” ? Why is lateral diffusion of integral proteins often appreciably faster in artificial lipid bilayers than in any of the natural membrane systems where extrinsic constraints are presumed to be absent? Diffusion is now generally referred to as cm2/second) for rhodopsin in disk membranes, band 3 in “rapid” (D 2 spherocytic erythrocytes, and several different proteins in blebbed membranes. It is commonly accepted that diffusion in this regime is limited soley by lipid “viscosity. ” However, dimyristoylphosphatidylcholine (DMPC) at 25°C is bound to be more viscous than disk membrane phospholipids, which contain on average about six cis double bonds per molecule. Yet purified bacteriorhodopsin diffuses about six times faster in fluid DMPC than does rhodopsin in the disk membrane (Peters and Cherry, 1982). Similarly, purified band 3 protein diffuses about sixfold faster in DMPC than it does in spherocytic erythrocytes (Chang et al., 1981; Koppel et al., 1981).’ ‘Other integral membrane proteins found to diffuse at 5 cm2/second when reconstituted into phospholipid bilayers include murine histocompatibility (H-2K) and vesicular stomatitis G proteins

PROTEIN DIFFUSION IN CELL MEMBRANES

33

Part of this disparity could easily derive from an increased bilayer viscosity due to the high protein content (ca. 50 wt%) of biological membranes relative to that of most reconstituted systems previously studied with FRAP. Cherry et al. (1977) found a 50-fold decrease in the rotational diffusion coefficient of bacteriorhodopsin in DMPC bilayers as the protein content was increased from 25 to 63 wt%. In analogy with the marked effects of dissolved soluble proteins on water viscosity (Fahey and Green, 1938; Treffers, 1940), they ascribed the drop in bacteriorhodopsin rotational rates to an increase in lipid viscosity due to the added protein. The observation of Jacobson et al. (1981) that lateral diffusion of lipid analogs is significantly faster in multibilayers constructed from fibroblast plasma membrane lipids than in the plasma membrane itself is also consistent with an effect of the protein on lipid viscosity, although the possibility of probe-protein binding also exists. The same argument applies to the finding that di-I diffusion in inner mitochondria1membrane preparations can be enhanced ca. three- to fourfold by dilution of the proteins with exogenously added phospholipid (Chazotte et al., 1983). Should integral proteins in reconstituted systems not be faithfully integrated into the bilayer then some enhancement of diffusion might ensue. As mentioned in Section II,E,3, lipid phase boundaries could restrict protein diffusion, and some NMR work on disk membrane lipids is consistent with the existence of two lipid domains, fluid and less fluid, with rhodopsin concentrated in the former (Brown et al., 1977). Adsorbed proteins like the GTP binding protein and phosphodiesterase in photoreceptor disk membrane, serum proteins on cultured cells and erythrocytes, or general nonfilamentous cytoplasmic proteins may also limit translational diffusion of integral proteins which protrude significantly into the aqueous phase. The effect of adsorbed BSA on diffusion coefficients cited in the previous section is in line with this idea. Finally, the intrinsically discrete (molecular) structure of membranes may help account for the discrepancy between the “viscous limit” in natural and model membranes. Saffman-Delbruck theory is a hydrodynamic model which treats the membrane as a featureless continuum characterized by a bulk viscosity. Yet we know that the third dimension of “two-dimensional” biomembranes is often asymmetric, with a unique lipid composition in each monolayer (see Smith ef ul., 1977, for evidence on photoreceptor disk membrane). Moreover, separate monolayers in these (Tanaka and Ohnichi, 1976; Schroeder, 1980) and artificial membranes (Seul et al., 1983) can apparently undergo independent phase changes. Diffusion in isotropic fluids may depend upon density fluctuations which open up local voids of free volume (Cohen and Turnbull, 1959). Is it (Cartwright et al., 1982); glycophorin (Vaz et al., 1981; Wu et al., 1981); the coat protein of M-13 phage (Smith ef 01.. 1979~);cytochrorne P-450 (Wu and Yang, 1980); and lipophilin (Derzko and Jacobson, 1978).

34

MICHAEL MCCLOSKEY AND MU-MING P O 0

possible that in anisotropic bilayers the diffusion of large transmembrane proteins is limited by the independent, and unequally probable occurrence of critically sized voids in opposing monolayers? It might be of interest to construct artificial bilayers both with and without pronounced transbilayer compositional differences and compare the diffusion rates of purified proteins therein. Related to the discontinuous structure of bilayers, we note that proteins can influence local lipid order, and that in principle this can lead to either attractive or repulsive interactions between the proteins (see references in Section III,A,2). Since these effects are most important at high protein concentrations, they must be considered when comparing diffusion rates in biological and model membranes. Such treatment is beyond our present scope however.

111. Implications in Membrane Biology A. PATTERNCREATION AND MAINTENANCE Living organisms are highly ordered and improbable arrangements of molecules, and only by constantly expending metabolic energy can they preserve this order. This is true for fluid “two-dimensional” membrane assemblies just as it is for three-dimensional structures like cells and tissues. Two striking examples where long-range membrane order is a prerequisite for biological function are the localization of ACh receptors at neuromuscular junctions in skeletal muscle and sodium channels at nodes of Ranvier in myelinated nerve fibers. Other examples include gap junctions, tight junctions, and epithelial cells with polarized lateral distributions of enzyme and transport activities. Short-range ordering of membrane components is important in the formation of nuclear pores, budding of membrane-enveloped viruses and adsorptive pinocytosis of polypeptide hormones through coated pits. In thylakoid membranes, sequestration of photosynthetic pigments in light harvesting and reaction center polypeptide complexes reflects an even finer pattern of order, probably crucial for efficient energy transfer. Diffusion is a direct manifestation of the Second Law, and superficially it is expected to counter the existence of organized features like those above. Certainly a cell must consume metabolic energy to contain potentially mobile proteins within functionally specialized regions of the membrane. The central question is how this is done. While many hypotheses have been advanced to explain immobilization and localization of membrane proteins, their success rate and predictive value have been unremarkable. In the following sections we briefly highlight some major concepts which have been entertained. Before doing so, it is worth noting that pattern creation and pattern stabilization are not necessarily inextricably coupled to one another. For instance, proteins could be actively

PROTEIN DIFFUSION IN CELL MEMBRANES

35

driven to their destination by linkage to a submembranous contractile apparatus, while they might be kept there by passive protein-protein interactions leading to the formation of immobile aggregates. While the two processes are intertwined in what follows, it will be useful to keep the distinction in mind. 1. Contractile Macromolecules The most popular models of pattern creation and maintenance invoke direct attachment of dispersed or patched proteins to the cytoskeleton or cytoskeletonassociated molecules. A previous section has covered some evidence for cytoskeletal participation in maintaining the localization of membrane proteins. ATP-driven, directed motion of the attached species to specialized zones on the cell surface, analogous to the sliding filament model of muscle contraction, has often been proposed to account for localizing movement. Because such a disproportionate amount of attention has already been paid to this subject, we defer to previous articles by de Petris (1977), Weatherbee (1981), Oliver and Berlin (1982), Levine and Willard (1983), and Heath (1983). 2. Protein Solubility and Aggregation Several theoretical papers on so-called “lipid mediated protein-protein interaction” have appeared (Gruler, 1975; Marcelja, 1976; Schroeder, 1977; Owicki and McConnell, 1979), one treating explicitly the creation of long-range order (Gershon, 1978). Experimental verification that lipid structural alterations can mediate membrane functions by promoting the association and dissociation of specific proteins was recently obtained by Siege1 et al. (1981). They found that in spinach chloroplast thylakoids the stoichiometric interaction of lightharvesting pigment-protein complexes with photosystem I1 (PS 11) reaction center complexes was progressively disrupted by successive addition of soybean lipids to the membrane, and that energy transfer to the PS I1 trap dropped off in parallel. Disruption of the laterally dispersed supramolecular structures was accompanied by formation of extensive lattices of the light-harvesting complex. Lipid-mediated protein-protein interactions, although potentially selective in nature, are really one manifestation of the overall problem of protein solubility within the bilayer. Two widely known structures to which protein insolubility may contribute are gap junctions and the purple membrane of Halobacterium. In both cases there is long-range membrane order due to laterally separated arrays to densely packed integral proteins, and in neither case are cytoskeletal components implicated as a stabilizing factor. For the gap junction, gross localization of individual connexons is a direct consequence of the intercellular interaction of complementary proteins in two contacting membranes. Whatever their mobility, once formed they cannot diffuse out of the cell-cell contact zone without first breaking apart. But that does not explain why connexons are often assembled in discrete, organized plaques within the contact zone. Perhaps the formation of

36

MICHAEL MCCLOSKEY AND MU-MING PO0

these plaques from individual connexons reflects a reduction in solubility over the separate connexon precursors, due for instance to proteolytic cleavage or other covalent modification. Or perhaps locally high concentrations of connexon proteins resulting from diffusion-mediated trapping exceed the solubility limit for that species and initiate precipitation. Crystallization of bacteriorhodpsin in vivo may be promoted by high bacteriorhodopsinconcentrationscoupled with the unique neutral lipid composition of Halobacterium membranes, which contain significant amounts of squalene. Pertinent to this speculation is the observation that increasing concentrations of cholesterol in fluid phoshatidylcholine bilayers cause progressive lateral segregation of purified bacteriorhodopsin (Cherry et al., 1980). Integral membrane proteins may be thought of as colloidal particles dispersed in a thin layer of fluid. Rules that govern the behavior of colloidal dispersions and the alteration of the behavior that results from specific molecular modification might well be applied to certain membrane situations. A balance of electrostatic repulsion between charged protein moieties in the aqueous phase and van der Waals attraction within the lipid matrix is likely to be an important factor for molecular interactions in the cell membrane, in particular, the formation of protein aggregates. Gingell (1976) and Gingell and Ginsberg (1978) have discussed interesting implications of colloidal theory in membrane biology. Rubin et al. (1981) used Smoluchowski colloidal aggregation theory to model the redistribution of integral membrane components which accompanies thylakoid membrane stacking and destacking, and derived a lateral diffusion coefficient for the PS I1 complex. 3. Bulk Membrane Flow Bulk membrane flow is an active process that could impart a motive force to membrane proteins (Bretscher, 1976; Harris, 1976). It was originally invoked to explain whole-cell locomotion and ligand-induced capping, but in principle is applicable to contact-induced redistribution. One theory (Bretscher, 1976) has it that flow is sustained by insertion of membrane lipid at one or more sources and simultaneous removal at a spatial remote sink. A molecular filter at the sink was originally postulated which would selectively retain proteins on the surface while passing the lipids through; the coated pit was later assigned this role (Bretscher et al., 1980). Rapidly mobile proteins will resist the flow by randomization against the concentrating effect of flow, but relatively immobile species like protein aggregates or “patches” will be swept up and concentrated at the sink, as observed during capping or surface receptors. There are some interesting experimental findings at least partially consistent with this hypothesis. For example, on different mammalian cell types several monovalent lipid “receptors” will patch upon treatment with multivalent ligands, and the patches migrate into a cap (Revesz and Greaves, 1975; Sedlacek

PROTEIN DIFFUSION IN CELL MEMBRANES

37

et al., 1976; Stem and Bretscher, 1979; Schroit and Pagano, 1981). As with capping of integral proteins, lipid capping is in some cases energy dependent and blocked by cytochalasin B. Using FRAP, Schroit and Pagano (1981) measured a lateral diffusion coefficient of 8.5 X cm*/second for a cappable lipid on murine lymphoma cells. Because the lipids in question cannot span the bilayer and since this rapid diffusion is not indicative of an interaction with transmembrane proteins, it is argued that direct cytoskeletal mediation of lipid capping is impossible. Some idea of why a monovalent lipid should even patch may help elucidate the mechanism of lipid capping. The classic observations of Ingram (1969) and Abercrombie et al. (1970) on the directed rearward motion of particles attached to the dorsal surfaces of advancing fibroblasts formed the primary experimental basis for the hypothesis of membrane flow. More recent kinetic experiments on the surface distribution of specific Golgi-processed proteins in locomoting fibroblasts are consonant with the earlier particle experiments, and have been interpreted as local insertion of Golgi-derived vesicles at the leading edge coupled with removal at several perinuclear sites (Bergmann et al., 1983). A few arguments could be raised against the efficiency of membrane flow as a mechanism of protein redistribution and localization. First, by itself it is not a very selective method for localizing proteins. Second, in cases where localization of a protein is undesirable, the protein must be capable of quite rapid translational diffusion (D > l o p 9 cm*/second) to remain dispersed under the influence of membrane flow (Bretscher, 1976). If one can believe the results from FRAP experiments very few plasma membrane proteins have such high mobility. Third, even if they do, some extra supposition must be added to the basic hypothesis to explain how specific proteins are diffusionally immobilized yet still susceptible to the flow. In summary, flow would provide a general impetus for directed migration of many membrane components. Given some selective molecular interaction at the sink this might offer a rate enhancement over other potential collection processes.

4. Localization by Electric Fields Exogenously applied electric fields can induce migration and segregation of cell surface components in many cultured cells (see review, Poo, 1981). There is a distinct possibility that endogeneous electric fields within tissue may serve to localize or segregate components in differentiated cells. There are two likely regions where this might occur. First, a steady potential across an organized layer of cells could segregate different components to two sides of the cell layer. In epithelial tissue, steady potential drops on the order of millivolts across the cell layer have been detected. This is within the same order of magnitude of steady potentials that were found to cause migration of cell surface components

38

MICHAEL MCCLOSKEY AND MU-MING PO0

in cultured cells. Second, high-frequency focal current occurring at localized regions in nervous systems, e.g., synaptic junctions, nodes of Ranvier of myelinated axons, and axonal branch points, may cause migration and localization of membrane receptors and ionic channels (see discussion in Fraser and Poo, 1982). The second possibility is of particular interest for neurobiology, since it suggests a possible cellular mechanism for use-dependent modulation of signal transmission, a key problem in understanding the plasticity of nervous systems. While direct in vivo experimental evidence is lacking, theoretical modeling of electromigration of membrane components in the presence of pulsed, focal fields suggests that under favorable conditions of high electrophoretic mobility to diffusion coefficient ratio (e.g., for protein aggregates), membrane components could be moved laterally by repetitive focal potentials (S. H. Young, personal communication). Lateral electrophoretic displacement of integral membrane components could also be induced by increasing their net electrical charge, for example by phosphorylation. In fact, a light activated protein kinase catalyzes phosphorylation of the light harvesting chlorophyll alb protein complex (LHC)in chloroplast thylakoids, and the phosphorylated LHC migrates out of appressed grana membranes and into the stroma exposed thylakoid regions. This event is believed to regulate the distribution of excitation energy between the two laterally separated photosystems, thus permitting a maximal quantum efficiency of photosynthesis under different light regimes (see Section III,D,2). Whether or not one accepts the specific models advanced to explain this phosporylation-induced redistribution, it is not hard to imagine that the high negative charge density within closely spaced grana stacks repels the negatively charged LHC. 5 . Tight Junctions in Segregation

Epithelial cell monolayers which carpet the lumen of organs like intestine, renal tubes, and pancreas perform a crucial homeostatic function in mediating the directional transport of nutritive substrates, ions, water, and digestive enzymes between the internal environment and the external world. Interstitial passage of these materials between apposed aqueous chambers is blocked to one degree or another by tight junctions, or zona occludens-specialized intercellular connections which seal the individual epithelial cells together into continuous sheets. Associated with this vectorial transport is a distinctly nonuniform lateral distribution of membrane components, which in the intestinal epithelial cell (IEC)is quite striking (Michell et al., 1976). Digestive enzymes and various sodiumsolute symport activities are concentrated in the lumenal membrane but nearly absent from the basolateral membrane. Transcellular sodium concentration gradients that drive nutrient absorption by these symport systems are created by Na-K-ATPase activity found almost exclusively in the basolateral membrane. Adenylate cyclase and receptors for various blood-borne stimuli like hormones

PROTEIN DIFFUSION IN CELL MEMBRANES

39

and cholinergic agonists are similarly restricted to the basolateral membrane. Some lipids are also asymmetrically distributed, with the cholesterol/phospholipid ratio of the lumenal membrane being twice that of the basolateral membrane. To the extent that plasma membrane proteins are capable of lateral motion, what prevents mixing of these various activities from bringing directional transport to a halt? Several groups have postulated that tight junctions preserve lateral asymmetry by blocking the translational diffusion of membrane components between lumenal and basolateral surfaces (Galli et a l . , 1976; Pisam and Ripoche, 1976; Parr and Kirby, 1979; Sang et a l . , 1979; Evans, 1980; Matsuura et a l . , 1982). Tight junctions disassemble in the absence of calcium, and isolation of epithelial cells dissociated by calcium chelators leads to randomization of polarized membrane topographies. For example, generally labeled glycoproteins and proteoglycans restricted to the lumenal face of frog urinary bladder epithelial cells become uniformly dispersed within 80 minutes of tight junction disruption (Pisam and Ripoche, 1976). Histocompatibility antigens confined to the basolateral membrane of mouse IEC also undergo complete redistribution after cell dissociation (Parr and Kirby, 1979). In pancreas and kidney epithelia the intramembranous particle (IMP) density observed with freeze-fracture electron microscopy is appreciably higher in the lateral than in the lumenal membranes, and in both cell types this polarity is lost concomitantly with cell isolation and tight junction disruption (Galli et a l . , 1976; Sang et al., 1979). Using freezefracture to follow the reappearance of IMP polarity in reassociating kidney cells Sang et al. (1979) observed that as soon as a single continuous string of junctional components had formed, differential IMP densities began to appear on opposite sides of the junction. Ziomeck et al. (1980) estimated the translational diffusion coefficients of leucine aminopeptidase (LAP) and alkaline phosphatase (AP) by measuring relaxation kinetics of the initially polarized distribution of these enzymes on freshly isolated mouse IEC. Although the relaxation rate was boosted by drugs which alter membrane potential and ATP levels, redistribution was apparently passive, with a half time of 20-30 minutes at 22°C. Because “though somewhat unraveled, substantial amounts of tight junctions are found on single cells and in cell pairs as late as 20 minutes after the IEC is isolated,” Ziomeck and associates deemed it unlikely that AP and LAP diffusion is restricted by tight junctions. Dragsten et al. (1981) showed that various fluorescent lectins bound to either the apical or basolateral surface of cultured kidney epithelial cell monolayers did not migrate to the opposite surface. However, using photobleaching they found the lectins to be essentially immobile (D < 10- l 2 cm2/second) along the apical membrane itself, and concluded that a barrier function for the tight junction need not be invoked to explain the polarized lectin distributions (see next section for alternative view). On the other hand, fluorescent antibody fragments bound to

40

MICHAEL MCCLOSKEY AND MU-MING Po0

the Na-K-ATPase in one of the same kidney cell lines are only 50% immobilized in either confluent or subconfluent cultures (Jesaitis and Yguerabide, 1981). In subconfluent cultures (junctions absent) the pump is uniformly distributed around the cells, while in confluent cultures (junctions present) it is restricted almost exclusively to the basolateral surface. Independent of the presence or absence of tight junctions, the mobile fraction diffuses at 5 X 10-lo cm2/ second. At this rate the mobile Na-K-ATPase molecules should be able to redistribute uniformly around the cell within an hour at most, provided they had a free path. To summarize, available evidence does not permit a definitive rejection or embracement of the hypothesis that tight junctions block lateral motion of integral membrane proteins; the possibility certainly remains viable for at least some proteins. In any case, diffusion barriers will only help in the maintainence of patterned topographies; localized insertion, direct cytoskeleton-mediated segregation, or some other mechanism must work in concert with the tight junction to form an initial pattern. Evans (1980) discusses evidence for some of these processes in his review on plasmalemma polarity in the liver epithelial cell. 6 . Entrapment in Extracellular Matrices In vivo, intestinal epithelial cells bear perhaps the thickest glycocalyx of any mammalian cell type. Smithson et al. (1981) have presented evidence that it impedes the diffusion of small molecules like sucrose, lactose, and oligopeptides to the plasmalemma enzymes which hydrolyze them. Thinner, but appreciable “fuzzy” coats also adorn the apical surfaces of other kinds of epithelial cells. In many instances this carbohydrate-rich layer is greatly depleted on the basolateral relative to the lumenal surface (Rambourg, 1971), and randomization of this pattern has been observed following tight junction disassembly and cell dissociation (Pisam and Ripoche, 1976). Lectins (MW lo4) are somewhat larger than sucrose, and one might expect their diffusion through the glycocalyx matrix to be significantly impaired; indeed, even unlabeled integral proteins which protrude into the external aqueous phase may suffer the same fate. Perhaps this helps explain the observations mentioned above that lectins bound to apical surfaces of epithelial cells are immobile, while Fab fragments bound to the Na-K-ATPase on basolureral surfaces have considerably greater lateral mobility. If this hypothesis is correct then the tight junction may block the lateral motion of lectin receptors in an indirect manner, by serving as a “dam” to retain certain glycocalyx constituents on the apical surfaces of epithelial cells.2 Leucine aminopeptidase and alkaline phosphatase are confined to the apical Qermane is the recent report by Rizki and Rizki (1983) that wheat germ agglutinin binding sites in the plasmalemma of Drosophila larval fat body cells are directly held in an asymmetric pericellular distribution by the adjacent basement membrane.

PROTEIN DIFFUSION IN CELL MEMBRANES

41

(lumenal) surface of IEC, yet Ziomeck et al. (1980) report fairly rapid diffusion cm2/ coefficients for these two proteins ( D values 0.5 X l o p 9 and 0.9 x second) upon cell dissociation. Perhaps this reflects glycocalyx thinning (redistribution) of the kind demonstrated by Pisam and Ripoche (1976). In addition, hyaluronidase treatment used in the isolation protocol may have trimmed the cell coat sufficiently to release severe diffusive constraints. 7. DifSusion-Mediated Trapping a. Evidence and Background. Diffusion-mediated trapping (DMT) is perhaps the simplest concept that embodies mechanisms for both development and maintenance of ordered biomembrane topographies. As a direct consequence of their Brownian motion, molecules will willy-nilly blunder into a trap region to which they chemically bind and simultaneously disappear from the surrounding membrane. In principle, the trap can be furnished by a neighboring cell or the host, and it may reside outside or inside the cytoplasm. If contributed by a touching cell it may consist of uniformly dispersed, mobile molecules having affinity for complementary species on the host cell. Upon comparison with other mechanisms its efficiency is fairly apparent; in the simplest case metabolic energy is spent only on the synthesis of mutually sticky proteins. DMT will clearly not explain all cases of pattern creation in biomembranes; for example, capping and various receptor redistributions which occur in chemotaxing, dividing, or phagocytosing blood cells occur too rapidly to be accounted for by diffusion alone (de Petris, 1977; Koppel et al., 1982; Oliver and Berlin, 1982). Other internally coordinated topographical rearrangements like those which occur during adsorptive endocytosis and virus budding may well follow the DMT route. We anticipate that molecular redistribution induced by membrane- membrane contact will prove to be a major biological application of DMT. Edwards and Frisch (1976) were the first to consider DMT in the control of cell surface topography. They attempted to account for ACh receptor turnover in neuromuscular synapses by postulating random insertion of newly synthesized ACh receptors in the extrajunctional region of the muscle followed by diffusion to the junction. In discussing the general problem of cell-cell adhesion Bell ( 1979) proposed that intercellular linkage of complementary receptors would result in the diffusion of more and more free receptors into the contact zone, thus rapidly increasing the strength of binding. Under experimental circumstances several contact-induced redistributions have been observed. When ferritin-conjugated Con A was used to agglutinate a mixture of pigeon erythrocytes and peripheral lymphocytes, the ferritin-Con A was found concentrated on the contacting regions of membrane and depleted elsewhere (Singer, 1976). Because both cell types bind Con A it was assumed that the tetravalent lectin acted as an intercellular bridging ligand, thus facilitating passive accumulation. Asialoglycoprotein receptors in the plasmalemma of rat hepatocytes undergo a strik-

42

MICHAEL MCCLOSKEY AND MU-MING PO0

ing accumulation in the zone of contact with surfaces of a synthetic galactosidecontaining polymer (Weigel, 1980). Galactose is the terminal hexose on asialoglycoproteins recognized by this receptor; replacement of galactose in the polymer with other sugar residues fails to induce receptor accumulation. According to Kampe and Peterson (1979), gene products of the major histocompatibility complex become localized in the contacting membranes of cytotoxic T lymphocyte-target cell conjugates. These are all suggestive pieces of evidence, but until recently neither the kinetics nor the mechanism of such processes had been investigated. Work in our laboratory has now provided several concrete examples of pure DMT, including the passive contact-induced accumulation of FcR-IgE complexes in the membrane of rat basophilic leukemia cells touching haptenated polyacrylamide beads (McCloskey and Poo, unpublished) as well as the concentration of both soybean and wheat germ agglutinin receptors in the contacting membranes of Xenopus embryonic muscle cell pairs in culture (Chow and Poo, 1982). Chao et al. (1981) have developed a quantitative model for molecular trapping by perfect sinks which relates the average lifetime of untrapped species to their diffusion coefficient and the area of membrane-membrane contact. Weaver (1983) has recently extended this theory to include imperfectly absorbing traps. b. ACh Receptor Localization. During synaptogenesis of the skeletal neuromuscular junction, ACh receptors over the surface of the embryonic muscle become highly concentrated at the subsynaptic muscle membrane. In Xenopus embryonic nerve and muscle culture, it was clearly demonstrated that contact by appropriate nerve processes can induce clustering of preexisting ACh receptors at the site of contact (Anderson and Cohen, 1977; Cohen and Weldon, 1980). The mechanism by which a nerve induces such a modulation of ACh receptor topography is unknown. Of various possibilities, the simplest explanation is that the ACh receptors, freely diffusing within the plane of the muscle membrane, are trapped at the site of nerve contact by binding to specific molecules associated with the nerve membrane or the intercellular matrix between the nerve terminal and the muscle. This diffusion-mediated trapping mechanism is plausible only if the diffusion rate of ACh receptors is rapid enough to account for the time-course of ACh receptor clustering both in vitro and in vivo, given the dimension of the developing muscle cells that diffusion must cover and the size of nerve-muscle contact. A rough estimate based on results obtained in culture (Cohen et al., 1979) and in intact tadpole (Chow, 1980) suggests that ACh receptors must diffuse with a D value of at least l o p 9cm2/second for the diffusion trap model to be plausible (Poo, 1982; Young and Poo, 1983). For the turnover rate of ACh receptors in mature muscle endplate to be explained by diffusion trapping a similar D value is also required (Edwards and Frisch, 1976). Early FRAP measurements of ACh receptor mobility in rat myotubes yielded a much smaller D value of 5 X 10- * cm2/second (Axelrod et a l . , 1976). Due to

PROTEIN DIFFUSION IN CELL MEMBRANES

43

the concern that FRAP is measuring diffusion of the complex of ACh receptor and its labeling ligand, a-bungarotoxin, rather than the native free receptor, an entirely different method, postinactivation recovery (PIR) was used to measure the diffusion of functional free ACh receptors in the muscle membrane. In this method, ACh receptors on muscle surface were locally inactivated by a pulse of a-bungarotoxin, a snake toxin that binds irreversibly to the ACh receptor. The recovery of ACh sensitivity at the site of inactivation was mapped by an iontophoretic method to reveal the lateral diffusion of the functional, toxin-free ACh receptors in the membrane. The D values obtained for both Xenopus culture preparations and intact developing muscle fibers of Xenopus tadpoles were found to be within the range of 1-4 X lop9cm2/second (Poo, 1982; Young and Poo, 1983). This diffusion rate is rapid enough to account for nerve-induced clustering of ACh receptors by a diffusion trap mechanism. What edge could the diffusion trap mechanism have over other conceivable mechanisms such as local insertion of ACh receptors at the nerve-muscle contact site together with selective removal of receptors from extrajunctional regions? Besides a possibly lower energy cost, diffusion-mediated trapping provides the simplest signaling mechanism for receptor localization. Teleologically speaking, the cell does not need to know where on the surface the nerve has made contact. All it has to do is to disperse ACh receptors on the surface; the receptor will be trapped wherever the contact happens to be made. This perhaps is one of the reasons why embryonic muscle puts a large quantity of ACh receptors into its plasma membrane before the nerve arrives (Blackshaw and Warner, 1976). c. Receptor-Mediated Endocytosis. Coated pits are small (50-200 nm) specializations of the eukaryotic plasmalemma which mediate the acquisition from extracellular fluids of a variety of nutritional, hormonal, immunological, enzymatic, and other macromolecular substances (Goldstein et d . , 1979). The prevailing conception is that these clathrin-coated depressions capture laterally diffusible ligand-receptor complexes from adjacent regions of the membrane and then pinch off intracellularly to form coated vesicles, which deliver the cargo to cytoplasmic compartments. The detailed dynamic events which occur between binding of ligands to their specific receptors and the appearance of clustered ligand-receptor complexes in coated pits have yet to be worked out; whether or not cells employ a simple strategy involving trapping of monomeric ligandreceptor complexes by preformed coated pits is currently unknown. Thus, some propose that ligand-induced receptor clustering provides a cue (e.g., membrane warping) that initiates clathrin assembly underneath the clusters (Roth and Woods, 1982). Notwithstanding the gaps in our knowledge regarding receptor-mediated endocytosis, it is possible to place certain restrictions on the lateral diffusion rates demanded by simple diffusion-mediated trapping of monomeric ligand-receptor complexes in preexisting coated pits. Assuming the coated pits to be perfect

44

MICHAEL MCCLOSKEY AND MU-MING Po0

sinks, Goldstein et al. (198 1) used published data on receptor densities, internalization rates, coated pit dimensions, copies per cell, and steady-state concentration of trapped vs free receptors to calculate the apparent two-dimensional rate constant for trapping of LDL receptors on a human fibroblast cell line. Upon comparison with the theoretical rate constant calculated from a previously measured D value of 2-3 x lo-" cm2/second (Barak and Webb, 1981) they concluded that if the foregoing mechanism does operate, it must do so at very close to the diffusion-limited rate. One can rationalize why a relatively small diffusion coefficient would suffice for the harvesting of LDL by considering the short distance involved. For human fibroblasts at 4°C the surface density of coated pits is about 0.58/pm2 (Orci et al., 1978), which gives an average interpit spacing of roughly 1.3 microns. In contrast, ACh receptors being trapped at the neuromuscular synapse must traverse the entire developing muscle, which may be hundreds of microns long. Because diffusion times depend on the square of the distances covered, the vastly different time frames of endocytosis and ACh receptor accumulation can be accounted for by the observed diffusion coefficients. Most indications are that insulin and epidermal growth factor receptors exist in a highly dispersed state prior to ligand binding. Ligand-induced clustering and internalization of EGF receptors is blocked at 4"C, and this could be due to a reduced lateral mobility of the receptors at 4°C if trapping of EGF were a diffusion limited process. Hillman and Schlessinger (1982) find only a threefold drop in the lateral D value of EGF-EGF receptor complexes over the span of 37 to 5°C and from this they estimate roughly the expected drop in encounter frequencies of bound receptors with coated pits. While they believe that harvesting of EGF proceeds by diffusion-mediated trapping, they conclude that the rates of visible patching and endocytosis of EGF receptor-EGF complexes are not determined by the lateral diffusion rates of EGF receptor-EGF complexes. A more conservative statement might be that low temperature inhibition is not a result of reduced diffusion rates; clearly, shifting the temperature can differentially affect the rate and equilibrium constants of separate elementary steps in a multistep sequence such that what was rate limiting no longer is. In the present situation this might arise if microtubule integrity were required for both visible patching and endocytosis. One might ask what advantages this diffusion-collection process offers. Would not the trapping and internalization of soluble ligands be faster if receptors were permanently glued to coated pits, thus bypassing steps dependent upon inexorably slow lateral diffusion? Alternatively, does spreading receptor sites uniformly over the plasma membrane improve the total catch of soluble ligands? Theoretical calculations (Berg and Purcell, 1977; DeLisi, 1981; Shoup and Szabo, 1982) lead to the prediction that for average cell dimensions the capture rate of soluble molecules is near maximal for only a few thousand sites per cell;

PROTEIN DIFFUSION IN CELL MEMBRANES

45

that is, the number of sites provided by coated pits alone should give efficient trapping. This does assume, however, that ligand binding by the cell is diffusion limited, and this assumption is not generally valid for all surface receptors (see Wank et al., 1983). It is known that several types of ligands can enter the cell through a single coated pit (Maxfield et al., 1978; Dickson et al., 1981; Willingham et al., 1981; Carpentier et al., 1982). Perhaps this sharing helps to explain the design of coated pit structure: There may be insufficient room inside a single coated pit to house several copies of all the different receptor types which interface with it. A multiple-user system may actually be simpler to coordinate than the evolutionary alternative of having many distinct kinds of coated pits, each with its own unique complement of receptors.

B. REACTIONS IN Two DIMENSIONS 1. Reduction of Dimensionality: Fact or Friction? a. Target Finding by Diffusion. Most membrane-mediated reactions fall into one of two categories: (1) Those involving water-soluble reactants and products which must find and depart the active site of a membrane-bound enzyme by some combination of two- and three-dimensional (2D and 3D) diffusion. (2) Reactions between more permanently attached molecules (e.g., integral enzymes, nascent polypeptides, and lipid-soluble isoprenoid coenzymes) where strict 2D interdiffusion governs collisional encounters. In 1968 Adam and Delbruck provided the first theoretical framework for analysis of membrane target finding via diffusion. They dealt specifically with the diffusion-limited trapping of water soluble molecules by a membrane-bound target which acts like a perfect sink (one must stretch the imagination a bit to picture enzymes and hormone receptors as infinitely absorbing, perpetually reusable traps). Based upon their calculations they suggested that a significant rate enhancement might accrue to membrane-mediated reactions if instead of finding the enzyme active site by purely 3D diffusive encounters a water-soluble substrate first adsorbed nonspecifically to the membrane and underwent a strictly 2D random walk until hitting the target. According to the theory this two-stage process, or “reduction of dimensionality,” during the diffusive search will increase the efficiency of target finding over that obtained in the absence of surface diffusion, provided that certain conditions are met. Namely, the adsorbed molecules must be sufficiently mobile (ratio of diffusion coefficients in 2D versus 3D: D,,lD3, L 0.01) and the ratio of diffusion space size to target size sufficiently small. For D,,lD,, 2 0.01 and an enzyme active site diameter of 20 A, the critical diffusion volume is about the size of a bacterium or a mitochondrion. For volumes much larger than this no advantage purportedly exists. Although in 1968 no lateral diffusion coefficients for any biomembrane components had yet been determined, Adam

46

MICHAEL MCCLOSKEY AND MU-MING PO0

and Delbruck nevertheless ventured that reduced diffusion times-presumably faster reactions-confer a selective advantage on the intracellular membranous organelles of eukaryotes; reduced diffusion times theoretically result from confining reactants within a subcritical volume and promotion of two-step capture by membranous subdivision of the aqueous space inside an organelle. Of course, all of this presupposes that the reactions of interest are diffusion-limited or would be without dimensionality reduction. Adam and Delbruck’s idea received an interesting twist when a more general analysis of membrane target finding by soluble reactants (Berg and Purcell, 1977; DeLisi and Wiegel, 1981) revealed that, in principle, purely 3D target searching can be more effective than one would suppose. Berg and Purcell reasoned that, due solely to statistical considerations, once a molecule has found the membrane it is guaranteed a period of diffusion near the surface, and thus has several more tries at bumping into the target before mixing with molecules out in solution. In effect, an inert membrane has its own “trapping” ability, causing the reactant to hop around awhile in a “quasi-2D” search. For reactants that do adsorb to the membrane a truly two-dimensional search can occur; whether this leads to an enhancement depends on the conditions described above, the adsorption energy (Berg and Purcell, 1977), rate of dissociation from the membrane (Richter and Eigen, 1974), and other difficulty estimated factors like the relative orientations of membrane-bound vs free reactants. b. Reaction Rates in 2 0 versus 3 0 . Anomalously rapid association of the lac repressor with its operator on DNA (Richter and Eigen, 1974; Schranner and Richter, 1978) and reduced transient times of some multienzyme complexes (Welch and Gaertner, 1975; Mosbach, 1976) provide the strongest support for rate enhancement by guided diffusion (dimensionality reduction). There is also some empirical evidence that under appropriate conditions the rate of reaction of aqueous with membrane-bound species can be enhanced by prior adsorption to the membrane (Overfield and Wraight, 1980). However, the common notion that reactions are necessarily “faster” in or on membranes than in the cytoplasmic aqueous phase has yet to secure a firm theoretical foundation. As discussed in the last section, the efficiency with which soluble ligands find membrane bound targets cannot be attributed simply to a reduction from 3D to 2D diffusion. Furthermore, even given a favorable ratio of 2D to 3D diffusion rates (say 0.01), it is not at all clear whether a significant theoretical rate advantage actually exists for reactions completely confined to the membrane plane. How does one compare the efficiency of a 2D reaction with one in 3D? What biologically relevant parameter, and what values of that parameter will actually indicate a selective advantage? It is our contention that if cells could think they would probably be most concerned with the total number of a given molecule they could make per unit time. A meaningful parameter to test the sweeping argument for reduction of

PROTEIN DIFFUSION IN CELL MEMBRANES

47

dimensionality might then be the net reaction rate ratio for equal numbers of reactant molecules in purely 2D and purely 3D reactions. One estimate of this ratio is obtained by comparing the collision frequencies of two reactants when confined to a membrane plane with that when contained within a 3D aqueous volume ensheathed by that membrane, i.e., the initial reaction rate of a totally diffusion-limited reaction (every encounter productive). According to Smoluchowski formalism, unlike in three dimensions, the rate “constant” for 2D diffusion-controlled reactions of the type A + B + C declines continuously with time (Emeis and Fehder, 1970; Naqvi, 1974) as it asymptotically approaches zero (Torney and McConnell, 1983). Hardt ( 1979) was apparently unaware of this in her comparison of reaction rates in one, two, and three dimensions, and her expressions for ID and 2D diffusion-limited rate constants are patently false. The only rigorous expressions suited to our calculation are those of Torney and McConnell (1983), and we employ these below. For simplicity, let us assume that the reaction involves two molecular species present in equal number; this is an arbitrary choice which should bias the result in favor of the membrane. To place the calculation into biological perspective, we take the reaction between cytochrome c and cytochrome oxidase in mitochrondria as an example. Estimates of cytochrome oxidase surface densities in mitochrondria range from about 1 per 50,000 to 1 per 590,000 A2 (Klingenberg, 1967; Hackenbrock and Hammon, 1975; Hochman et al., 1982), giving a total number of oxidase between 0.2 and 1.7 X lo4 for a spherical mitoplast of radius 0.5 km. Since the ratio of cytochrome c to the oxidase varies from about 0.8 to 1.7 (Capaldi, 1982), our assumption of equal numbers of each reactant is reasonable. The encounter radius is assumed to be 20 A. Making simplifying assumptions about the lattice spacing, for times greater than a small fraction of a second we find the ratio of diffusion-controlled reaction rates in 2D vs 3D to be:

RJR, = 83(D,/D,)/ln(AD,t) where A is of the order of l O I 4 cm-,. Recalling that membrane proteins in general diffuse two to three orders of magnitude more slowly than soluble proteins in buffers, and that the empirically determined diffusion rates of cytochrome c and the oxidase (see Section 111,B,2) are no exception, this maximum possible “enhancement” could actually be an impediment, especially at long times. In the above case, even if one inserts the greatest D, yet observed for a membrane protein (5 X 10W9 cm2/second) and a reasonable D, of lo-’ cm2/ second, the membrane-mediated reaction is still appreciably slower than the bulk phase reaction (R2/R, = 0.3). Notice that we have compared the hypothetical reaction rates for a maximum ratio of volume to surface area (spherical vesicle). Some cellular organelles contain flattened membranes, e.g., chloroplast grana stacks, mitochondria1

48

MICHAEL MCCLOSKEY AND MU-MING PO0

cristae, and pancake-shaped Golgi stacks. Reactions in the aqueous space between the appressed membranes can be thought of as approaching the extreme of reaction within a plane. Drawing on the previous result, since diffusion within these spaces might be as fast as in free solution, we suggest that diffusion-limited reactions between aqueous and membranous reactants may be enhanced somewhat therein. c. Anisotropic Rotation and Orientation Constraints. In contrast with diffusion-limited reactions between small molecules, it is unlikely that very many reactions between macromolecules are controlled simply by collision frequencies of the reactants. After all, large molecules with one active site are not uniformly reactive over their surface and not all collisions are expected to result in a favorable alignment of the reactive sites. For example, we can infer from results of Schmitz and Schurr (1972) that the surface domains involved in electron transfer from cytochrome c to peroxidase probably occupy 10% (or less) of the total surface area of both proteins. Similarly, only 7-15% of the surface area per subunit is inaccessible to solvent in protein complexes such as the insulin dimer, the ap dimer of hemoglobin, and the trysin-BPTI complex (Chothia and Janin, 1975). That this heterogeneous reactivity can have a large influence on the rates of diffusion-limited biochemical reactions is indicated by the very low probability (steric) factor (1.8 X for head-tail joining of bacteriophage T4D (Aksiyote-Benbasat and Bloomfield, 198 1). All indications are that large amplitude rotational motion of integral membrane proteins is confined to an axis normal to the membrane plane. This restricted rotation permanently aligns the reactive groups in a way that is impossible for soluble macromolecules, and one might expect this orientation to provide some rate advantage in reactions between large proteins. A rigorous theoretical treatment of the effect of orientation constraints on reaction rates is beyond the scope of this article, but a semiquantitative calculation based upon probability arguments is informative. Assume the two reacting proteins to be spherical in shape with radii a, and a, and each to have a circular reactive patch of radius b, or b, on its surface. For simplicity, let a1 = a, and b , = b,. Assume that reaction ensues when there is overlap of these sites during a collision. For proteins in aqueous solution the probability that a collision will result in overlap of reactive sites is the product of fractional areas occupied by the sites on each molecule. Next suppose the same proteins are immersed in a bilayer and constrained to rotate about an axis normal to the membrane-such that the reactive sites are correctly positioned to interact. In this arrangement the appropriate fractional area is the ratio of target area to the area swept out by the target in rotating through 360" about the axis normal to the membrane, and the probability of a productive collision is the product of this ratio for both proteins. The net ratio of probabilities in two dimensions to three dimensions is then, roughly

PROTEIN DIFFUSION IN CELL MEMBRANES

49

P21P, = (a/b)2 If we go back to the example of cytochrome c reacting with peroxidase this probability ratio is estimated to be about 10 (Schmitz and Schurr, 1972). This means that the actual rate ratio for 2D versus 3D might be noticeably greater than predicted solely on the basis of collision frequencies, and that reduction of the dimensionality of rotational diffusion (3D to 1D) may provide more of a rate advantage than reduction of dimensionality of translational diffusion (3D to 2D), at least when realistic lateral diffusion coefficients are used in the calculation. This simple (simplistic?) treatment considers only the probability of reacting upon the first contact, and neglects the possibility that an ineffective collision might be remedied by rotational searching within the encounter complex (Solc and Stockmayer, 1973; Simmons, 1975). d. Conclusions. We have shown in the above two sections that when reasonable diffusion rates and reactant concentrations are considered, standard collision frequency calculations do not substantiate the general conception that reactions are “faster” in membranes than in the aqueous phase-r that the speed of strictly membrane-bound reactions confers a selective advantage on intracellular membraneous organelles. However, we call attention to two previously unexplored aspects of dimensionality reduction. First, bulk phase reactions or membranous target searching could proceed faster within the intermembranous aqueous space of flattened intracellular membranes than in unbounded systems, without actual partitioning of the reactants into or on the membrane. Some potential candidates are plastocyanin-mediated electron transfer in thylakoid lumens, cytochrome c redox reactions along mitrochondrial cristae, and sorting of soluble proteins within Golgi stacks. Second, because integral proteins are held by the membrane in a reactive orientation, the probability that a given collision will be productive should be greater than for similar proteins in aqueous solution. Just what the magnitude of this effect actually is and whether it is great enough to override the characteristically slow lateral and rotational diffusion of membrane proteins in conferring a net 3D to 2D advantage remains to be seen.

2. Electron Transport a. Respiration. The mechanistic pathway of intra- and intermolecular electron transfer along the respiratory sequence is a challenging problem with several unresolved aspects. Exactly how these redox reactions drive ATP synthesis also remains a mystery. Those who subscribe to the chemiosmotic hypothesis cannot agree on whether a proton concentration gradient itself or the associated transmembrane electrical potential gradient is most important, and some nonsubscribers think that local (lateral) proton gradients and not transmembrane gradients are the most immediate link to ATP synthesis (Storey and Lee, 1981; Wikstrom, 1981; Williams, 1981; Haines, 1983). At the lowest level of resolution one is

50

MICHAEL MCCLOSKEY AND MU-MING PO0

concerned with the spatial organization of separate electron carriers within the inner mitochondrial membrane, and with its time dependence. Early concepts which equated the chemical sequence (decreasing redox potential) of individual components with a static physical sequence, or “respiratory chain” (Lehninger, 1959), have gradually given way to more dynamic models which picture at least some components as randomly distributed within the membrane plane and without any permanent binding between reaction partners. At the extreme of dynamicism are those who contend that the entire respiratory sequence, consisting of dehydrogenases, coenzyme Q, the cytochrome bc, complex, cytochrome c, and cytochrome oxidase (Fig. l), is completely jumbled and that electron transfer rates are determined solely by collision frequencies, and hence the lateral diffusion rates of separate molecules within the inner mitochondrial membrane (Hackenbrock, 1981). We devote extra space to this topic not only because of its paramount importance to living organisms, but also to document our assertion that meaningful interpretation of diffusion measurements is virtually impossible without a detailed knowledge of the biochemistry and ultrastructure of biomembranes. Early evidence on chain dynamics. One of the initial findings which weakened the physical “chain” concept was that complete electron transfer from NADH or succinate to oxygen could be produced in systems reconstructed from isolated complexes I-IV, cytochrome c , and ubiquinone (Hatefi, 1968). Klingenberg was the first to point out that individual cytochromes and flavoproteins (NADH and succinate dehydrogenases) do not occur in a 1:l mole ratio within inner membranes, and also that significant variation in the overall mole ratios exists between different tissues in one organism and between different species as well. Many investigators seem to consider this strong evidence against the existence of a specific supramolecular complex between different proteins of the respiratory sequence. Communication between physically discrete chains fed by succinate or NADH would have to occur at an early step, since succinate will reduce all the cytochrome be, molecules. The Q-pool behavior reported by Kroger and Klingenberg (1973a,b) was interpreted in just this way, that is, shuttling of electrons between separate chains by laterally mobile coenzyme Q. Relative lateral mobility of at least some portion of cytochrome c was implicated by the experiments of Wohlrab (1970), who found that membrane-bound cytochrome c can mediate the equilibration of iron oxidation states in cytochrome oxidase molecules located within separate “chains. ” Modulation of electron transport by exogenous lipids. In a series of ingenious experiments Hackenbrock and associates have manipulated the spacing between integral proteins of inner mitochondrial membrane preparations and studied the effects on rates of electron transport between separate portions of the respiratory sequence (Schneider et al., 1980). They found that when increasing numbers of sonicated lipid (soybean) vesicles are fused with inner membrane vesicles

PROTEIN DIFFUSION IN CELL MEMBRANES

--+C

SUCCINATE

51

Yt

FUMARATE

FIG. 1 . Mitochondria1 electron transport chain: Complex I , NADH dehydrogenase; Complex 11, succinate dehydrogenase; Complex 111, cytochrome bc, complex; Complex IV, cytochrome au3 complex (cytochrome oxidase); UQ, ubiquinone; Cyt C, cytochrome c.

(mitoplasts) from rat liver mitochondria the average spacing between intramembranous particles (IMPs) increases. Even though the separate complexes (I-IV) retain the same or show increased specific activities, electron transport from NADH and succinate to the bc, complex drops off appreciably with increased spacing between the IMPs. Schneider etal. (1980) concluded that electron transport between these molecules is a diffusion-controlled process, the increased interparticle distances reducing collision frequencies. Diffusion-mediated may be a more apt descriptor since all reactions (both diffusion- and activation energycontrolled) go slower when the reactant concentration is dropped. But even this conclusion is tempered by the results of another incorporation experiment where the average interparticle spacing was progressively reduced by cholesterol-induced lateral segregation of the inner membrane proteins (Schneider et al., 1982). Although cholesterol incorporation prevented a drop in rates of electron transport from NADH and succinate to complex 111, the rates never climbed above control levels-even though the average spacing within IMP rich domains became much shorter than in unmodified mitoplasts. If random encounters were the whole story, then shorter distances should generate faster reactions just as longer paths lead to slower reactions-assuming that electron transport remains diffusion-mediated. C. R. Hackenbrock (personal communication) suggests that imperfect lateral segregation led to entrainment of crucial electron carrier proteins in the proteinpoor domains. Another possibility is that ubiquinone remains uniformly dispersed as the proteins segregate, thus reducing the ratio of this coenzyme to complexes I-IV within the protein-rich region. Although these studies have correlated increased areas per IMP with slower reactions, they have not established a cause and effect relationship between the

52

MICHAEL MCCLOSKEY AND MU-MING PO0

two. Since respiration rates can only be lowered from native values by lipid incorporation, it is entirely possible that exogenous soybean lipids (which are structurally distinct from endogenous rat mitochondria1 lipids) cause progressive disruption of protein-protein interactions normally required for efficient electron transport. Such interactions may be more sensitive to the lipid environment than is tacitly assumed in most studies with reconstructed lipid-protein systems. This was vividly demonstrated by Siege1et al. (1981) in completely analogous vesicle fusion experiments with chloroplast thylakoid membranes, where dilution of endogenous lipids with soybean lecithin led to progressive detachment of light harvesting chlorophyll-protein complexes from photosystem I1 reaction centers and consequent decrease in energy transfer efficiency. Consideration of lipidmediated effects on the organization and function of membrane proteins is imperative in studying modified or reconstituted systems. Lateral difusion measurements. In contrast to NADH and succinate oxidase activities, Schneider et al. (1980) found little decrease in duroquinol oxidase activity as the amount of incorporated lipid was increased. (Duroquinol oxidase activity is electron transport from a water soluble hydroquinone to the bc, complex through cytochrome c and the oxidase to oxygen.) To rationalize this they suggested that cytochrome c either diffused very rapidly between physically separate bc, and aa3 complexes or that the three components diffused together as a unit. Since subsequent work showed that the oxidase and bc, complexes patch independently when inner membranes are treated with a mixture of anti-bc, and anti-aa, antibodies the former hypothesis was favored. To test this possibility Gupte er al. ( 1 983) measured diffusion of fluorescein-labeled cytochrome c on large inner membrane preparations formed by calcium induced fusion of mitoplasts. They found that the apparent lateral D value is markedly dependent on ionic strength: in 0.3 mM Hepes D = 4.0 X l o - " cm2/second; in 10 mM KP, D = 2.5 X 10WLocm*/second; in 25 mM KP, D = 2 X cm2/second. Electron transport from succinate or duroquinol to oxygen increased moderately on going from 0.3 mM Hepes to 10 mM KP, (factor of 1.2-1.5). Hochman et al. (1982, 1983) also estimated D values for rhodamine labeled cytochrome c over the surface of mitoplasts from megamitochondria of cuprizone-fed mice. Although the effect of ionic strength was not reported, the number obtained in 8 mM Hepes (1.6 X 10- lo cm2/second) is essentially the same as what Gupte et al. (1983) obtained in 10 mM KP,. In both experiments bound cytochrome c was present at 10 times the level found in intact mitochondria (endogenous cytochrome c was first depleted). One wonders if only 10%of the exogenous cytochrome c occupies sites that its natural counterpart does in a whole mitochondrion; appreciably faster or slower diffusion of this fraction might go unnoticed. It is interesting that while both groups find about the same D value at low ionic strength, one contends that lateral diffusion of cytochrome c is far too slow (at

PROTEIN DIFFUSION IN CELL MEMBRANES

53

least an order of magnitude) to account for observed reaction rates, while the other group favors the notion that electron transfer by cytochrome c is diffusion mediated. Calculations based upon mean free paths (Hochman et a l . , 1982) and Hardt’s equations (LeMasters et a l . , submitted) have been used to argue against or for the sufficiency of these measured D values in supporting completely diffusion-mediated electron transport. When we repeat these calculations using Adam-Delbmck (1968) or Torney-McConnell (1983) equations we find that lop9 cm2/second is apparently too slow to account for some of the more rapid rates of respiration reported for intact mitochondria (Moreadith and Jacobus, 1982), but perhaps sufficient to explain the intermediate to lower rates. However, this conclusion rests precariously upon input parameters such as the reaction radius, oxidase surface density (monomer-dimer?), steric factors, etc., and until these are defined more accurately the significance of this arithmetic remains obscure. The rationale for focusing on cytochrome c mobility was that motion of the integral complexes appeared too slow to support a collisional mechanism of electron transport between any parts of the sequence. Sowers and Hackenbrock (1981) were the first to measure lateral D values for these proteins; using a combination of postfield relaxation and freeze-fracture electron microscopy to quantitate IMP distributions they estimated an average diffusion coefficient of 8.3 X 10- l o cm2/second for all the IMPS in spherical mitoplasts from rat liver mitochondria. The same group later found a D value of 4.0 X cm2/second for complexes 111 and IV using photobleaching methods (personal communication). Hochman et al. (1983) also used FRAP to quantitate diffusion of complex IV in “megamitoplasts” from cuprizone fed mice and obtained a diffusion coefficient of 1.0 X 1 O - I o cm2/second. Again, whether any of these rates is sufficient to support a purely collisional mechanism of electron transport (i.e., 1/2 turnover per hit) depends rather critically on the distances involved, which are still uncertain (see Hackenbrock and Hammon, 1975). Suffice it to say that all these diffusion coefficients pertain to mitoplasts which have been rendered spherical by incubation in low osmolarity buffers, and that a substantial portion of the matrix proteins (240%) is lost during this process (Caplan and Greenwalt, 1966; C. R. Hackenbrock, personal communication). When one considers that the mitochondria1matrix contains between 0.35 and 1.05 g protein/ml (Capaldi, 1982) and that this range encompasses the composition of protein crystals, the loss of some 50% may lead to a major experimental artifact. Others (Srere, 1982; Capaldi, 1982) have already intimated that the motion and organization of integral proteins of the inner membrane may be affected by this dense matrix. One reason that cytochrome c lateral diffusion rates at physiological ionic strengths have not been reported is that the labeled protein does not stick to membranes very well in more concentrated buffers. The ionic strength inside a mitochondrion is likely to be greater than that of 0.2-25 mM buffers, and as

54

MICHAEL MCCLOSKEY AND MU-MING PO0

Gupte et al. (1983) point out, three-dimensional diffusion of cytochrome c within the intermembrane space of intact mitochondria “may be an important component of diffusion-mediated electron transfer. This is an interesting possibility since the translational diffusion rates of small proteins like cytochrome c in water are nearly three orders of magnitude greater than the largest rates measured for cytochrome c on membranes (Barisas et al., 1979; Cadman et al., 1981). While mitoplasts do carry out electron transport at low ionic strength (where all cytochrome c is membrane-bound), it should be noted that this is not necessarily at the maximal rates observed for whole mitochondria, and is therefore a weak argument against the contribution of aqueous phase diffusion to electron transport. However, it is not apparent from our crudely modified Berg-Purcell type calculations that even at cm*/second purely 3D diffusion would generate sufficient collision frequencies to explain the high electron transfer rates reported for rat heart mitochondria (Moreadith and Jacobus, 1982). This very approximate calculation ignores geometrical effects of the cristae folds; it is tempting to speculate that because these flattened membranes nearly confine the path of aqueous cytochrome c to lie in a plane that electron transport between bc, and aa3 enjoys the full kinetic benefit of 2D diffusion that Adam and Delbruck originally referred to, albeit for different reasons (Section III,B, 1). Biochemical/ kinetic evidence on be,-c-aa3 sequence. There are several other lines of evidence regarding the organization and dynamics of the bc,-c-aa, segment. It is now well documented that essentially the same amino acid residues on cytochrome c are involved in binding to not only the oxidase and reductase but also to other proteins such as cytochrome b, and cytochrome c peroxidase (Ferguson-Miller et af., 1978; Speck et al., 1979; Stonehuerner et al., 1979; Poulos and Kraut, 1980; Rieder and Bosshard, 1980; Azzi et al., 1982). These are positively charged lysines which encircle the heme crevice and form a complementary template to negatively charged carboxylate groups surrounding the exposed heme edge in the interacting proteins. The fact that bc, and au3 complexes bind to essentially the same residues on cytochrome c has been used to argue that the latter must diffuse laterally between these two components. However, limited rotation of cytochrome c within a ternary complex could also be a workable model-a model in concert with the observation that cytochrome c covalently bound to either the reductase or the oxidase is still capable of mediating electron transfer (Erecinska et al., 1980; Waring et al., 1980). In support of this scheme, the reduced form of cytochrome c dissociates from purified complex 111at least 10 times faster than the oxidized form. Ferguson-Miller er al. (1978) also envisage simultaneous complexation of cytochrome c by both the oxidase and reductase as being consistent with their kinetic results. Relevant to this model, Hackenbrock and Hammon (1975) made the interesting observation that monospecific antibodies against purified cytochrome oxidase will displace most of the endogenous cytochrome c from inner membranes, ”

PROTEIN DIFFUSION IN CELL MEMBRANES

55

The kinetics of electron transfer from cytochrome c through the oxidase to oxygen are complex, and exhibit two or three phases (Thompson et al., 1982); with purified oxidase and cytochrome c the polarographically determined rate of oxygen reduction is more than an order of magnitude faster than the net rate of appearance of oxidized cytochrome c as determined spectoscopically (FergusonMiller et al., 1978; Smith et al., 1979a). This coupled with other kinetic data has been interpreted to mean that the initial rapid phase of the reaction is due to multiple turnovers of bound cytochrome c prior to dissociation from the oxidase. In these experiments small molecular weight reductants like ascorbate and TMPD were used, and these have more ready access to the heme center than the physiological electron donor. Nevertheless, one is forced to seriously consider the possibility that once bound to the oxidase, cytochrome c turns over several times before the two become translationally separated. As far back as 1956 Chance and Williams discussed the general possibility that limited rotation or oscillation of separate components within a multiprotein complex could explain observed respiratory electron transfer rates. In fact, based upon spectroscopically measured kinetics in submitochondrial particles, Nicholls (1976) concluded that at least 80% of the endogenous cytochrome c in mitochondria is directly bound to the oxidase. Rotational diffusion measurements. Consistent with this idea, Dixit et al. (1982) found that a phosphorescent derivative of cytochrome c rotates at the same slow rate as cytochrome oxidase when bound to mitochondria1 membranes ( T= ~ 300 ksec). In this experiment the phosphorescent analog was substituted for native cytochrome c in an equivalent quantity to that present in vivo, although as with the fluorescent derivatives, subphysiological ionic strengths were necessary to prevent desorption. Saffman and Delbruck (1975) predicted that rotational diffusion coefficients of membrane proteins should be quite sensitive to molecular radii (D,cx r - 2 ) , and experimental findings are in accord with this (Cherry, 1979; Hughes et a l . , 1982). Since cytochrome c is much smaller than the oxidase it might be expected to rotate appreciably faster if it were nonspecifically bound as a monomer to the negatively charged lipid bilayer. The coincident rates may be fortuitous but they are suggestive of complexation between cytochrome c and the oxidase at low ionic strength. The data do not appear to exclude the possibility mentioned above of rapid, yet low amplitude rotation. Finally, we note that rotation rates of cytochrome oxidase reconstituted in lipid vesicles are not influenced by the simultaneous incorporation of the reductase (complex 111) (Kawato et a f . , 1981). Combining this with the relative independence of duroquinol oxidase activity and intramembranous particle density which Schneider et al. (1980) observed in lipid incorporation studies, Kawato and associates conclude that rapid lateral diffusion of cytochrome c between spatially discrete bc, and aa3 complexes is more likely than formation of a ternary complex of the three molecules. There is one obvious reason, however, why this reconstituted system may simulate with low fidelity the interactions within a

56

MICHAEL MCCLOSKEY AND MU-MING PO0

mitochondria1 inner membrane, namely, the lipid milieu is much different. As emphasized in a preceding paragraph, if lipid-mediated protein-protein interactions are involved in the maintenance of an intact, functional structure, then the arbitrary substitution of a synthetic lipid (fluid or otherwise) for the complex blend indigenous to a cellular organelle could disrupt that structure. Summary. While a large body of evidence weighs against the existence of a single static electron transport chain, much uncertainty remains regarding the dynamic structural organization of the terminal segment of the sequence. Several simplified or reconstituted systems which are amenable to direct measurements of protein diffusion have yielded interesting clues about the role of lateral diffusion in respiratory electron transfer. Nevertheless, for reasons made clear in the above, the relevance of measurements on these systems to the intact mitochondrion remains an open issue. b. Photosynthesis. Current dogma regarding the mechanism of photosynthetic electron transport and phosphorylation in green membranes is embodied by the so called Z-scheme (Fig. 2), which has at its heart the concept of noncyclic electron transport. According to this picture two independent photochemical reactions activated by light of different wavelengths operate in series to drive the endergonic transfer of electrons from water to ferredoxin and then NADP. The initial photoact (680 nm) generates an oxidant capable of stripping electrons from water and a weak reductant which donates electrons to a sequence of carriers including plastoquinone, the cytochromeflb6 complex, and plasto-

Q

r

Lo

REDUCTION POTENTIAL

-

WATER

FIG. 2. Photosynthetic electron transport chain: PSI, photosystem I (trap WOO); PSII, photosystem I1 (trap P680); PQ, plastoquinone; Cyt blf, cytochrome bdfcomplex; PC,plastocyanin; Fd, ferredoxin; FNR, ferredoxin-NADP oxidoreductase.

PROTEIN DIFFUSION IN CELL MEMBRANES

57

cyanin. The second photoact (700 nm) produces a weak oxidant that accepts electrons from plastocyanin and a strong reductant capable of reducing ferredoxin, a high potential Fe-S protein. Energy released during the exergonic transport of electrons from Q to plastocyanin is somehow utilized to make ATP, which along with NADPH is consumed during fixation of atmospheric carbon dioxide in dark reactions (Calvin cycle). Photophosphorylation of ADP is also coupled to some cyclic electron flow driven only by the second photoact. The two photochemical events occur in chemically distinct photosystems, which are now known to consist of unique proteins containing one reaction center chlorophyll a and many (order of lo2) highly oriented antenna chlorophylls which transfer excitation energy to the reaction center, or trap. Photosystem I1 (trap P680) mediates water oxidation while photosystem I (trap P700) reduces ferredoxin in both cyclic and noncyclic electron transport. The bulk of the chlorophyll a and essentially all chlorophyll b is contained in a third pigment-protein complex which conducts no known photochemistry. As its name suggests, a primary function of this light harvesting complex (LHC) is to transfer excitation energy to the photochemically active pigment-protein complexes. It is now quite certain that the lateral distribution of various components of the thylakoid membrane is markedly nonuniform. Thus, the ATPase and photosystem I are totally excluded from grana partitions (the appressed regions of grana stacks), while photosystem 113, cytochrome b,,,, and the LHC are highly concentrated there (Anderson and Anderson, 1982, and references therein; Usharani et al., 1983). The antibody accessibility of ferredoxin-NADP reductase (Jennings et al., 1979) is consistent with an earlier suggestion (Berzborn, 1969) that it is located primarily in stroma-exposed regions of the thylakoid. On the other hand, the cytochrome f / b 6 complex appears to be partitioned equally between appressed and exposed thylakoid regions (Cox and Anderson, 1981; Anderson, 1982). Average diameters of grana stacks range from 0.3 to 0.5 p,m, and stroma lamellae can be somewhat longer. Thus, relatively long-range lateral separation of the two photosystems presents an interesting biophysical problem for the conventional Z-scheme, namely, is lateral diffusion of the intermediate electron carriers fast enough to cover such distances within their turnover times? Anderson and Anderson (1982) suggest that plastoquinone and plastocyanin should be capable of mediating long distance electron transport by a diffusional mechanism. Indeed, if one simplifies the argument by assuming that all the pho3Several groups have presented spectroscopic evidence for two distinct types of PS 11: PS I1 a and PS I1 (3 (Melis and Hohman, 1976; Horton and Croze, 1979; Thielen and Van Gorkom, 1981). PS I1 p (the minor fraction of PS 11) is supposed to have fewer antenna chlorophyll a molecules associated with it than PS I1 a.Anderson and Melis (1983) now report that PS I1 a is found exclusively in grana partitions while PS I1 p is restricted to stroma exposed thylakloids.

58

MICHAEL MCCLOSKEY AND MU-MING PO0

tosystem I is located in margins of the grana thylakoids, then a back of the envelope calculation is possible. With PS I1 uniformly dispersed in the appressed membrane, the minimum (average) distance a quinone must diffuse during one turnover (20 msec) can be taken as twice the radius of the stack, or 0.4 pm. This would require a minimum lateral diffusion coefficient of 2 x 10V8 cm*/second, which is about as fast as any lipid analogs have been observed to diffuse in biomembranes (at physiological temperatures). In fact, the requirements on diffusion rates are more stringent than we assume in the above. First, the actual diffusion paths may be substantially greater, since a significant portion of PS I is located not in the margins and end membranes but in the fret membranes. Second, plastoquinone (PQ) may diffuse much slower than proponents of the diffusion mechanism will allow. Nobody has ever directly measured the lateral diffusion rate of this rather long lipid molecule nor has anyone yet proven that it is not bound to a carrier protein. Third, turnover times for other redox components that are candidates for lateral electron transport, e.g., plastocyanin, are appreciably shorter than 20 msec (Haehnel et al., 1980), and this demands even faster d i f f u ~ i o n Clearly, .~ a rigorous analysis depends critically on the values of several variables, none of which is known with tremendous certainty. Even though the immediate prospect of incorporating diffusional parameters into a realistic model for photosynthetic electron transport is small, our current knowledge of the lateral segregation of the two photosystems dictates that noncyclic electron transport via the conventional Z-scheme must involve comparatively long-range lateral diffusion of plastocyanin and perhaps the cytochrome b/f complex. Arnon and associates have recently challenged the notion of noncyclic electron flow in higher plants. According to the current Z-scheme PS I1 cannot directly photoreduce ferredoxin; only PS I is supposed to be a strong enough reductant to accomplish this. Using different inhibitors of electron transport that block at plastoquinone, Arnon et al. (1981) find that exogenous ferredoxin is directly reduced even when PS I is poisoned. Further, using the aqueous polymer twophase partition method to isolate vesicles highly enriched in PS 11, direct electron transfer from water to NADP is again observed (Arnon et al., 1983). The authors postulate that ferrodoxin is used as an electron acceptor by both PS I and PS 11, and that two separate photoacts involving PS 11 drive electron flow through a circuit involving plastoquinone, cytochrome blf and PS I1 associated cytochrome 4Diffusion in the aqueous phase near the membrane may play an important role in energy transducing membranes (see Section III,B,I). Cox and Anderson (1981) attempt to account for the rapid cm2/second) in the aqueous turnover of plastocyanin by postulating that it diffuses rapidly (ca. medium within the intrathylakoid space. Based upon their FRAP studies, Gupte et al. (1983) advance a similar argument for rapid diffusion of cytochrome c in the aqueous phase between inner and outer mitochondria1 membranes.

PROTEIN DIFFUSION IN CELL MEMBRANES

59

b,,,. While this hypothesis places less stringent requirements on the lateral mobility of plastocyanin and plastoquinone, Arnon believes that these species must still carry “a trickle” of electrons between the two photosystems, for regulatory purposes. Therefore, both of the major working hypotheses on photosynthetic electron transport appear to require a corrolary hypothesis which invokes relatively long-range lateral diffusion of some component(s) of the electron transport chain. An interesting ramification of Amon’s proposal stems from the apparently nonuniform lateral distribution of ferredoxin-NADP reductase. Ferredoxin reduced by PS 11 must diffuse from the appressed to the stroma exposed thylakoids before it can reduce NADP. Ferredoxin is water soluble, and easily lost from chloroplast membranes during isolation procedures. What percentage of the ferredoxin in an intact chloroplast is actually adsorbed to the membrane and what percentage is free in the stroma is not known. Thus, whether it would have to diffuse laterally on the membrane or free in solution is a moot point; the latter would seem inimical to maximal production of NADPH, but ferredoxin does participate in other reactions involving soluble phase reactants (e.g., thioredoxin). Relevant to the mode of linkage of ferredoxin to the membrane it is interesting to note that Wagner et al. (1982) find a 40-fold reduction in the rotational rate of membrane bound ferredoxin-NADP reductase upon addition of ferredoxin. Because ferredoxin is small (MW 12,000) it is not expected to reduce the rotational diffusion rate of the reductase (MW 35-40,000) nearly so much upon binding; the authors speculate that ferredoxin mediates the formation of a trimolecular complex between PS I, ferredoxin, and the reductase.

C. BIOLOGICAL SIGNALING AT

THE

MEMBRANE

Signal is defined here as a piece of information transferred between a cell and its environment. In the following discussion we consider the possible relevance of lateral diffusion to both the reception and propagation of signals at the plasma membrane. Some topics, for example the putative role of hormone receptor mobility in cyclic AMP-mediated responses have already received considerable attention in other places. However, two subjects from neurobiology and immunology involve a previously unemphasized concept, namely, diffusion-mediated signaling within the plane of the membrane. 1. Transmembrane Signaling

a. Cyclic AMP-Mediated Hormonal Response. The literature is replete with references to the possible significance of lateral mobility in cyclic AMP (CAMP)mediated hormonal response, and the subject needs little introduction here. Adenylate cyclase systems consist of at least three physically separable membrane proteins which interact upon hormone or neurotransmitter binding to catalyze

60

MICHAEL MCCLOSKEY AND MU-MING Po0

synthesis of CAMP:the catalytic unit, the hormone (or neurotransmitter)receptor, and the GTP binding protein. The bulk of evidence is most easily rationalized with a “mobile receptor” hypothesis, which postulates that collisional encounters between independently diffusing catalytic units and receptor- hormone complexes produce the catalytically active state. Two observations supply the best evidence for this hypothesis. (1) Hormone stimulated cyclase activity can be restored in heterokaryons, one partner of which lacks functional catalytic units and the other of which lacks functional hormone receptors (Orly and Schramm, 1978; references in Schulster, 1979); (2) after irreversible inactivation of 93% of the p-adrenoreceptors in turkey erythrocytes, the remaining 7% can activate all the catalytic units, though at a slower rate (Tolkovsky and Levitsky, 1978). It was originally envisaged that receptor-ligand complexes interact directly with catalytic units, and discussion centered on the relative lateral mobility of just these two components. The situation assumes interesting complexity now that the existence of a completely separate GTP binding unit (G unit) is fully recognized. Rodbell (1980, 1981) discusses evidence from target size analysis, and proposes that in some cell types the G units and receptors are copolymerized into oligomeric structures in the membrane; according to the model, hormone or neurotransmitter binding triggers dispersal and frees the regulatory (G) and receptor units for interaction with the catalytic unit. This hypothesis may be rather speculative at present, but we find the results of a recent FRAP study in striking concert with one of its predictions. Henis and Elson (1981) observed that on Chang human liver cells both antagonist-labeled and (one infers) unliganded preceptors are present as visually discernible, immobile clusters over the cell surface. The small mobile fraction has a rapid lateral D value of 1.4 X cm*/second at 23°C. Agonist binding causes slow (minutes) dispersal and mobilization of receptors without affecting lipid diffusion rates. On the other hand, the time course of mobilization is much slower than that of cyclase activation (seconds) but roughly parallels the kinetics of receptor loss and enzyme desensitization. The inescapable conclusion is that ‘‘adenylate cyclase activation by preceptors does not require macroscopic lateral mobility of the majority of preceptors”-at least in Chang human liver cells. Others have also speculated that the cAMP response to hormones might be regulated through control over the lateral mobility of cyclase components. Hirata and Axelrod (1980) for example, suggest that specific ligands can induce local enzymatic modification of the lipid milieu, and this is supposed to enhance cyclase activity by increasing the frequency of encounters between receptors and catalytic units, e.g., because of greater lipid fluidity. The finding that microtubule disassembly can increase hormonally stimulated cAMP synthesis is consonant with cytoskeletal regulation of the mobility of cyclase components (Insel and Kennedy, 1978; Rudolph et al., 1979). Rasenick et al. (1981) observed that

PROTEIN DIFFUSION IN CELL MEMBRANES

61

colchicine and vinblastine not only enhanced the G unit mediated activation of cyclase activity in rat synaptic membrane preparations, but also loosened the connection of G units to the membrane sufficiently that a sizable portion could be removed after one rinse in the centrifuge. They concluded that “the ability of the G unit to diffuse laterally in the membrane is a limiting factor in cyclase activation. ” Cis-unsaturated but not saturated fatty acids also enhanced cyclase activity, but without knocking G units off the membrane. Combining these results with the ability of cis but not trans-unsaturated or saturated fatty acids to block capping in leukocytes (Klausner et al., 1980), a new interpretation may be suggested for the initial finding (Hanski et af., 1979) that the “fluidizing agent” cis-vaccenic acid speeds agonist-induced activation in membrane fragments from turkey erythrocytes. It was found that the “rate constant” for hormone activation of cyclase increased linearly by 2O-fold over less than a 2-fold variation in the bilayer microviscosity; perhaps this was not due so much to increased lipid fluidity as to decreased interaction of some cyclase component with the cytoskeleton. We can only conclude that future studies which directly probe the time dependence of rotational and translational diffusion of individual cyclase components during activation will help firm our grasp on the actual mechanism of this physiologically important response. Resolution of the coupling mechanism also hinges on more detailed structural and biochemical knowledge of the noncovalently linked components. Development of fluorescent reporters with specific affinity for the individual proteins (see, e.g., Haley et al., 1983) seems to be the primary hurdle both for photobleaching and for resonance energy transfer experiments aimed at measuring distances between the individual sites. b. Leukocyte Degranufation. Receptor clustering and dispersal have frequently been considered candidates for the initial actuating event in several different immunological, hormonal, and neurological responses. Speculation on this topic has been fueled by the following kinds of evidence: bivalent antibody induced patching and capping in lymphocytes, PMNs, and fibroblasts (dePetris, 1977); polyclonal stimulation of lymphocytes by mitogenic lectins, bacterial lipopolysaccharides, and other potentially multivalent ligands; the dependence of antibody response on antigen valency and state of aggregation (Dintzis et al., 1976, 1982, and references therein); clustering of informational macromolecules prior to internalization (Goldstein et al., 1979; Schlessinger, 1979);the ability of bivalent polyclonal antibodies against the insulin receptor to mimic certain biological responses produced by insulin (Kahn, 1979; Baldwin et al., 1980); potentiation of the response to low concentrations of insulin or EGF by bivalent antibodies against these hormones (Shechter et af., 1979a,b); stimulation of testosterone synthesis in Leydig cells by bivalent antibodies against the lutropin receptor (Podesta et af., 1983); data referred to above on the adenylate cyclase system. Should it ever be demonstrated conclusively that passive receptor ag-

62

MICHAEL MCCLOSKEY AND MU-MING PO0

gregation (or dispersal) really is an essential activating step, then local translational (and rotational) diffusion of membrane proteins will be directly implicated in the elicitation of physiological response. Perhaps the least equivocal example where clustering of specific cell surface receptors is a sufficient condition for cell triggering is the antigen-stimulated degranulation of mast cells and basophils. These are highly specialized leukocytes (basophils) and connective tissue cells (mast cells) that form an integral part of the immediate allergic response to soluble stimuli. On their surface are high affinity, monovalent receptors for the Fc stem of IgE (Mendoza and Metzger, 1976); although serum IgE levels are very low (nanomolar) IgE Fc receptors are normally at least partially (20-100%) saturated in vivo (Malveaux et al., 1978). The interior of these cells is packed with membrane surrounded granules containing histamine (and/or serotonin), heparin, and protein. Interaction of multivalent antigens (like ragweed proteins) with cell borne IgE causes granules to fuse with each other and with the plasmalemma, thus releasing histamine into the bloodstream. Several biochemical reactions reportedly accompany (or precede) degranulation, including phosphatidylserine decarboxylation, N-methylation of phosphatidylethanolamine, phospholipase-mediated arachidonic acid release, prostaglandin and leukotriene synthesis, CAMPproduction, activation of two protein kinase isozymes, and calcium influx (reviewed by Ishizaka, 1982); but the primary event seems to be antigen-induced bridging of IgE Fc, receptor complexes (reviewed by Kagey-Sobotka et a l . , 1982). In fact, neither IgE nor cross-linking antigen is required. Bivalent antibodies against the purified receptor will cause degranulation (Ishizaka et al., 1971; Isersky et a l . , 1978), as will bivalent antibodies against IgE when applied to IgE-sensitized cells (Ishizaka and Ishizaka, 1978); chemically cross-linked dimers of IgE in the absence of antigen will also trigger mast cells (Segal et al., 1977). Degranulation does not require global changes in the membrane, since locally applied stimuli cause degranulation directly beneath the stimulus (Lawson et al., 1978; Diamant et al., 1970). Extensive mathematical modeling of this system has been performed, and if the competing process of desensitization is allowed for the calculations predict with reasonable accuracy dose-response curves for histamine release in the presence of structurally well-defined bivalent haptens (DeLisi, 1979; Dembo et al., 1979a,b; Chabay et al., 1980). Using a range (100-fold) of experimentally measured rate constants to work backwards from the model and predict a diffusion coefficient for the IgE Fc, receptor complex, DeLisi (1979) obtains numbers to 10- cm2/second, a region which straddles the FRAPin the range of determined value of 2 X 10- l o cm2/second (Schlessinger et al., 1976). c. Cell-Cell Communication via Diffusion-Mediated Trapping. There are some recent observations on basophil degranulation which may have far-reaching biological implications in terms of generalized signalling mechanisms in

PROTEIN DIFFUSION IN CELL MEMBRANES

63

cell-cell communica$on. While it is certain that relatively long-lived and thermodynamically stable pairwise interactions between Fc, receptors are sufficient to generate the primary biochemical/biophysical events leading to degranulation of mast cells and basophils, they may not be necessary. Rat basophilic leukemia (RBL) cells bearing anti-dinitrophenyl IgE will release serotonin upon contact with either fluid or solid phospholipid vesicles containing a monovalent and most probably monomeric DNP lipid hapten (Balakrishnan et al., 1982). Assuming that dimerization or oligomerization of Fc, receptors remains an obligatory step in stimulation by the lipid vesicles, one would like to know what drives receptor aggregation in the absence of cross-linking reagents. Balakrishnan and co-workers suggest, among other possibilities, that diffusion-mediated localization and concentration of receptors in the vesicle-cell contact zone may contribute. We have observed substantial and rapid accumulation of fluorescent, cell-bound antiDNP IgE at the region of contact with DNP-derivatized polyacrylamide beads, even in the presence of 30 mM NaN,. Provided that a prior equilibrium exists between monomeric and “clustered” IgE receptors, locally high IgE receptor concentrations may shift the equilibrium in favor of “clusters” and thus initiate degranulation. If specific receptor clustering is a widespread and basic signaling device then diffusion-mediated trapping may offer a general mechanism whereby two mobile, independently diffusing components on opposing cells can participate in cell-cell communication. Bell (1979) proposed an analogous model to explain the interaction of histocompatibility-restricted cytotoxic T lymphocytes with virally infected target cells.5 In a provocative report Kampe and Peterson (1979) showed that histocompatibility antigens actually do localize in the contact zone between killer T cells and their targets, although the mechanism of localization was not investigated. One wonders if lymphocyte triggering by antigen presenting macrophages is mediated in part by passive contact-induced accumulation of antigen and receptors in the contact zone. Dintzis and associates (1976, 1982) find that the primary antibody response to T cell-independent antigenic polymers depends critically on the number of haptens per polymer chain, and propose that formation of a specific, well-defined cluster of lymphocyte receptors, or immunon, is an obligatory step in triggering antibody synthesis. They suggest that cell-cell contact between lymphocytes and “helper” cells with clustered surface-bound antigen could also generate the putative immunons; based upon the results of Balakrishnan e? al. (1982) it may be simpler to postulate that in the case of T cell5Local as well as long-range diffusion is implicated in this and other cell-cell interactions where specific receptors must bind to complementary species on an adjacent cell. As shown in studies on macrophage phagocytosis (McConnell, 1979; Lewis ei al., 1980) this is most evident at low receptor/ ligand densities, where the probability of molecular bond formation as well as the maximally attainable numb& of bonds can be enhanced by relative lateral diffusion of the interacting components.

64

MICHAEL MCCLOSKEY AND MU-MING PO0

dependent antigens, monomeric antigenic fragments on the macrophage surface (bound to Ir gene products?) induce clustering of lymphocyte receptors by a combination of diffusion-mediated trapping and Le Chatelier’s principle. We stress that diffusion trapping is a passive process which in the above applications can only promote clustering if the intercellularly bridged receptors remain mobile. d. Contact-Induced Immobilization. At the opposite extreme, it seems that complete immobilization of a freely diffusing molecule by binding to an immobile species on a neighboring cell could also pass information through the membrane, and the response might be faster in this case. Imagine, for example, that the cytoplasmic concentration of some key molecule was kept relatively constant and spatially uniform by continual collision dependent activation of a plasmalemma enzyme like adenylate cyclase. The sudden and local immobilization of activator molecules would cause a quick and initially localized drop in the cytoplasmic level of this key molecule, provided the enzyme was relatively immobile. Immobilization of naturally occurring membrane-associated inhibitory substances, e.g., the peripheral protein inhibitor of mitochondria1 ATPase (Emster et a l . , 1979), GABA-modulin (Wise et al., 1983), certain glycosaminoglycans which directly inhibit adenylate cyclase (Cutler, 1982), would also polarize the cell interior. While this group of molecules does not actually possess all the requisite attributes, it does serve to illustrate the general idea of how local, contact-induced freezing of molecular motion might work to promote information transfer between cells.

2. Information Flow in the Membrane Plane a. Counting Synaptic Contacts. “Signaling” usually connotes specifically the transmembrane delivery of messages. In principle, however, two cells can communicate without information ever leaving the membrane plane, through membrane reorganization mediated by lateral diffusion of membrane components. Formation of synaptic connections provides an interesting example. In a developing nervous system each postsynaptic cell receives a certain number of nerve terminals, usually more than the number left in a mature nervous system. In neonatal skeletal muscle fibers, each fiber is innervated by several separate axons (Brown et al., 1976); as maturation proceeds all but one synapse is eliminated through synaptic competition. Concomitantly with establishment of synaptic transmission, receptivity of the muscle surface to further innervation is lost. Similar events occur during synaptognesis between neurons (Purves and Lichtman, 1980). It appears that muscle and nerve cells can somehow count the number of cell-cell contacts both in the embryonic multi-innervated state and in the mature state. After a preset number of synaptic contacts is made, the extrasynaptic region of the membrane loses receptivity to further contact. One simple mechanism that would account for counting is based upon diffusion-mediated

PROTEIN DIFFUSION IN CELL MEMBRANES

65

trapping. If the initial recognition/adhesion between neurite and postsynaptic cell is mediated by some mobile species of recognition/adhesion molecule that accumulates and becomes trapped at the developing synapse, then concurrent depletion of the molecule from the extrasynaptic membrane will eventually prevent the recognition/adhesion with more neurites. Thus, by regulating the amount of the hypothetical recognition/adhesion molecule present on the cell surface (e.g., by controlling its rate of synthesis and degradation), the cell could determine the allowed number of contacts without the need for any signal as to where on the surface the contacts have been made. The receptivity of embryonic and denervated adult skeletal muscle fibers to innervation correlates well with the presence of high ACh receptor concentrations. Might not the ACh receptor itself be a recognition/adhesion mediating glycoprotein, in accordance with the above hypothesis? If so, nerve-induced accumulation of ACh receptors would then explain three separate phenomena of neuromuscular interactions: contact stabilization, localized channel activity, and control over the number of synapses (signaling). After maturation contacts may be stabilized by accessory interactions, such as with components of the basement membrane (Sanes et al., 1978). b. Immune Adherence. Macrophage Fc, receptors bind to IgG molecules that are present as opsonin on various phagocytosable particles and promote the attachment and engulfment of these foreign or altered-self targets. From a detailed mathematical analysis of macrophage-target binding kinetics (Lewis er al., 1980) coupled with independent observations from quantitative electron microscopic (Petty et al., 1981) and biochemical (Mellman et al., 1981) studies the following picture has emerged: Fc, receptors are irreversibly removed from the cell surface by the phagocytic internalization of plasma membrane. A reservoir of membrane is made available to the surface during this phagocytic challenge such that the total plasmalemma area remains nearly constant even though 50% or more may be internalized during a typical experiment. Fc, receptors remaining on the cell surface apparently undergo complete redistribution after particle uptake and reinsertion of fresh, receptor deficient membrane but prior to subsequent particle binding and uptake (this is true for phagocytosis of IgG opsonized 1 pm lipid vesicles by murine macrophages). That is, they rapidly diffuse into the receptor deficient patches. It may be that “down regulation” of IgG Fc receptors in response to phagocytosis is part of an appestat mechanism for reticuloendothelial cells, which by controlling the strength of immune adherence prevents phagocytes from overtaxing their finite digestive capacity for particles like opsonized Streptococcus and senescent erythrocytes. Perhaps rapid lateral redistribution of Fc, receptors permits fine tuning of the response by ensuring a uniform-rather than locally concentrated4istribution of attachment sites. Depending on the size and position of newly incorporated membrane patches, a nonuniform topography due to

66

MICHAEL MCCLOSKEY AND MU-MING PO0

IgG Fc receptor immobility could adversely prolong particle adherence. Although there is independent evidence of membrane recycling in macrophages (Muller er al., 1980), we know of no reports defining the size of newly added vesicles. Should their diameter be significant relative to that of a bacterium (say 0.4 Fm) then the above receptor dilution hypothesis is reasonable. Michl et al. (1 983) find that unligated Fc, receptors on murine macrophages diffuse at faster than 2 X l o p 9 cm*/second at 37”C, a rate compatible with rapid redistribution and two-dimensional “signaling. ”

D. SELF-ASSEMBLY AND SORTING One of the basic problems in cell biology is to understand the underlying biogenic processes which give rise to differentiated structures so characteristic of living organisms. Self-assembly and sorting of cellular components, both in the membrane and the cytoplasm, are key elements in these processes. Many membrane components are multimeric structures which are most likely self-assembled in the membrane from individual subunits. ,Some pertinent examples are cytochrome oxidase, photosynthetic protein-pigment complexes, Na-K-ATPase, prothrombinase, the membrane attack complex of complement, acetylcholine receptors, and gap junction connexons. Although very little is known about the detailed molecular events leading to self-assembly of multisubunit integral proteins, it is axiomatic that short-range translational diffusion of the individual subunits is mandatory. On the other hand, sorting mechanisms must somehow overcome the countervailing tendency of diffusion to randomize molecular distributions. A complex example of this interplay is found in eukaryotes, where the Golgi stacks are actively engaged in sorting of many membrane components. While a continual influx of new (from ER) and recycled membrane impinges on the Golgi, a melange of newly made and recycled proteins is constantly sorted and re-routed to the ER, lysosomes, secretory vesicles, or plasmalemma. At one step or another, these proteins must undergo long-range lateral separation in the membrane plane. While there is no lack of hypotheses pertaining to sorting within the Golgi complex, in reality next to nothing is known regarding the molecular mechanisms employed. For reviews see Tartakoff (1980), Rothmann (1981), Rothmann et al. (1981) and Olden ef al. (1982). Somewhat more is known about the following three examples, which should highlight the complexity of sorting problems, and also shed some light on the importance of protein diffusion in same. 1. Virus Budding One of the most dramatic cases of membrane sorting occurs during the budding of a membrane-enveloped virus from an infected cell. The viral envelope

PROTEIN DIFFUSION IN CELL MEMBRANES

67

forms from preexisting host cell membrane, yet it contains at most only trace amounts of host-determined proteins, the bulk (99+ %) of the envelope protein content being contributed by one or a few virally coded transmembranous glycoproteins (for review see Lenard, 1978). Enclosed by the envelope is an often highly symmetrical nucleocapsid consisting usually of single stranded RNA in close association with one or a few virus-determined nucleocapsid-, or N-proteins. With a few exceptions (the togaviruses), there is also a nonglycosylated matrix- or M-protein situated in the tight space between nucleocapsid and membrane. Both the M- and N-proteins are synthesized on “soluble” polyribosomes and reach the plasmalemma independently of the viral gl ycoproteins, which take the membrane-bound route via the rough endoplasmic reticulum and Golgi complex. In the final stage of assembly a preformed nucleocapsid, integral glycoproteins, and M-protein coalesce at the plasma membrane (usually) and a new viral particle buds off. In a positive or negative sense, protein diffusion is involved in at least three of the problems which attend virus budding: First, what generates the forces required to mechanically distort the membrane of a forming virion? Second, how are host cell membrane proteins selectively excuded from the budding virus? Third, how are virus glycoproteins selectively included in the budding virion? The magnitude of this “sorting” problem is not trivial; on the basis of published data on the protein content of typical plasma membranes and of vesicular stomatitis virus (VSV) envelopes (Cartwright et al., 1972), one can estimate that the surface density of host cell proteins can be less than one-tenth that in the precursor plasma membrane. Over the years several models for budding of membrane-enveloped viruses have been proposed (for references see Johnson et al., 1981). Regarding the mechanical properties of membranes, Evans and Buxbaum (1981) have shown that given sufficiently strong membrane-membrane attraction erythrocytes will passively and nearly totally engulf membranous vesicles as large as 1-3 pm. Willison et al. (1971) found that 0.12-pm latex spheres are partially swallowed by tomato protoplasts, and suggested that adhesive forces were sufficient to warp the plasmalemma passively in the early stages of endocytosis. Likewise, Haywood ( 1975) has demonstrated that liposomal membranes containing glycolipids which bind to Sendai virus will adhere to and tightly encase the virion. If a viral nucleocapsid had affinity for viral membrane glycoproteins and/or M-protein it would presumably nucleate a similar process when it made contact with the membrane; diffusion-mediated trapping or two-dimensional crystallization of M and/or glycoproteins around the “nucleus” would provide a ready explanation for exclusion of host cell proteins. That such simple mechanisms do not operate in VSV is evidenced by the presence of several temperature-sensitive (ts) mutants which form virus particles severely deficient in or entirely lacking nucleocapsids (Schnitzer and Lodish, 1979). Other ts mutants of VSV lack the membranous viral glycoprotein (Deutsch, 1976; Little and Huang, 1977) but

68

MICHAEL MCCLOSKEY AND MU-MING PO0

contain normal amounts of M-protein and nucleocapsid. This coupled with the significant (nearly twofold) variation in glycoprotein concentration that can occur in viral envelopes (Lazarowitz et al., 1973) is inconsistent with (1) pure steric hindrance due to tight packing of viral glycoproteins as the mechanism for exclusion of host membrane proteins (in those viruses containing M-proteins); (2) a cooperative aggregation of viral glycoproteins elicited by high glycoprotein concentrations. Apparently, no M-protein-deficient mutants with the ability to bud have been observed (Lenard, 1978); in this regard, it is interesting that Mprotein is not detectable in the virulent, nonbudding form of the subacute sclerosing panencephalitis strain of measles virus but is present in wild type and nonvirulent SSPE strains, both of which form buds (Lin and Thormar, 1980). This is suggestive evidence that M-protein is a central organizer essential for budding and possibly also for excluding host proteins from the viral envelope. Perhaps Mproteins are analogous to clathrin, which polymerizes into virus-sized baskets which bind to the plasmalemma, and which may form part of the structure within coated pits which discriminates between trappable and untrappable membrane components (e.g., ligated and unligated hormone receptors). The minimum structuraUcompositiona1requirements for budding have yet to be determined. While diffusion-mediated trapping of viral glycoproteins is probably not a universal mechanism for exclusion of host proteins, it may well induce segregation of glycoproteins into virally molded domains. For VSV and other M-protein-containing viruses there is a large excess of viral glycoprotein in the plasmalemma during budding, and for wild-type VSV infections of baby hamster kidney cells and chick embryo fibroblasts this component is diffusely distributed and largely mobile (75%), with a moderate diffusion coefficient of ca. 6 X 10- lo cm*/second (Reidler et al., 1981; Johnson et al., 1981). Qualitatively at least, these results are consistent with a diffusion trap mechanism. Reidler et al. (198 1) have argued from their photobleaching experiments with wild type and M-altered VSV mutants that specific complexation of M-protein with the glycoprotein is a rate-limiting step in budding of wild-type VSV particles but that reduced interaction of the altered M-protein with nucleocapsid becomes rate limiting in the mutants. The nature and extent of this putative interaction were unspecified. In contrast to VSV, Sindbis virus (SV) glycoproteins in the host plasmalemma are mostly immobile soon after infection starts (Johnson et al., 1981). This correlates with electron microscopic evidence that SV nucleocapsids become attached to intracellular membranes prior to reaching the plasmalemma (Johnson and Schlesinger, 1980; Gottlieb et al., 1979; Birdwell et al., 1973), perhaps implying that for this and other togaviruses, redistribution of viral glycoproteins from a diffuse to an aggregated state begins intracellularly. Since M-proteins are absent from these viruses, a simpler assembly may occur than for viruses con-

PROTEIN DIFFUSION IN CELL MEMBRANES

taining M-proteins-involving glycoproteins.

69

direct binding between nucleocapsid and

2. Thylakoid Stacking Thylakoid membranes in chloroplasts of higher plants and green algae are differentiated into grana stacks and stroma lamellae. As noted previously (Section III,B,2), the lateral distribution of several components of the photosynthetic apparatus is markedly nonuniform along the plane of the thylakoid membrane. While the true physiological significance of this topography is unknown, current thinking is dominated by the notion that it facilitates adaptation of the photosynthetic apparatus to different qualities and intensities of light. Thus, shade plants typically have higher ratios of grana to stroma lamellae than sun plants (Anderson et al., 1973); the ratio of stroma exposed to appressed thylakoid membrane area is also greater in algae cultivated in low light intensity than in algae grown in bright light (Reger and Kraus, 1970). Chloroplasts from higher plants and green algae respond within minutes to shifting light regimes (e.g., 650 vs 710 nm) such that maximal quantum yields are sustained (Bonaventura and Meyers, 1969; Murata, 1969; Punnet, 1971); this “State” transition is accompanied by partial thylakoid destacking and lateral redistribution of the light harvesting chlorophyll a / b protein complex (LHC) (Punnet, 1971; Bennoun and Jupin, 1974; Biggins, 1982; Staehelin et al., 1982). The transition is thought to reflect a balancing of the distribution of excitation energy between the two photosystems such that rates of noncyclic electron transport are optimized (Wang and Meyers, 1974; Butler, 1978). It is well established that red light stimulates phosphorylation of the LHC, and recent experiments have confirmed that the phosphorylated LHC moves out of PS I1 territory and into PS I enriched domains (Kyle et al., 1983; Staehelin et al., 1982). A cogent argument has been advanced that the redox state of plastoquinone regulates the kinase responsible for this phosphorylation, and in this way controls the relative rates of excitation in PS I and PS I1 traps (Bennet et al., 1980; Horton and Black, 1980). For full details on this model and references to the literature consult the review by Haworth et al. (1982). Passive diffusion-mediated trapping of the LHC appears to direct the stacking process per se, and once local environments are formed they may exclude other species due to steric constraints or electrostatic repulsion. In greening plastids the appearance of LHC polypeptides parallels the formation of stacked membranes (Armond et al., 1976; Davis et al., 1976; Argyroudi-Akoyunoglou and Akoyunoglou, 1977); mutants deficient in the LHC have no grana stacks (Anderson, 1975; Arntzen et al., 1976; Burke et al., 1979). Purified LHC undergoes reversible cation-mediated aggregation to form two-dimensional sheets which superficially mimic the grana/stroma system seen in intact chloroplasts (Mullet

70

MICHAEL MCCLOSKEY AND MU-MING P O 0

and Arntzen, 1980). As noted above, partial unstacking accompanies the phosphorylation-induced migration of LHC into stroma-exposed membranes. In virro, chloroplasts lacking the envelope undergo reversible cation-dependent stacking and destacking concomitantly with lateral redistribution of the LHC and other components (Izawa and Good, 1966; Goodenough and Staehelin, 1971; Ojakian and Satir, 1974; Staehelin, 1976). A small trypsin releasable peptide from the hydrophilic surface of the LHC is an absolute necessity for salt induced stacking and lateral segregation of the LHC (Mullet and Artnzen, 1980; Staehelin et al., 1980). From the time evolution of energy transfer between PS I1 units during artificial stacking, Rubin et af. (1981) calculate lateral D values in the range of 3.1 X to 1.9 X 10-l2 cm2/second (30 to 10°C) for PS I1 complexes. These numbers seem surprisingly low for proteins embedded in membranes composed of such highly unsaturated lipids and lacking obvious extrinsic constraints. Since the calculations are based on the assumption of a diffusion-limited process, the low D values may reflect some other rate-limiting step; blind application of the 3D Smoluchowski formulation to diffusion-limited processes in two dimensions is also perhaps unjustified (see Section III,B,l). In any case, it is probably only a matter of time before FRAP is used to directly measure the lateral diffusion coefficients of thylakoid components, not only to study the stacking process but also to investigate the role of lateral diffusion in photosynthetic electron transport (Section III,B ,2). The intrinsic fluorescence of the three major pigment-protein complexes should facilitate this work, and also preclude interfering effects of large labeling reagents like antibody fragments. No one knows the exact chemical nature of the "trap" for LHC, and speculation runs the gamut from specific polypeptide-polypeptide binding (lock and key) to nonspecific hydophobic interaction made possible by cation screening of the negative charges on opposing membranes. Barber (1980) and Rubin et al. (1981) present mathematical models of the stacking/segregation phenomenon based upon colloidal aggregation theory. PS I1 in the appressed regions is bound to the LHC, and it is not hard to imagine that LHC acts as an indirect trap for PS 11. Likewise, it is easy to conceive of the large extrinsic portion (CF,) of the ATPase as too big to fit into the appressed regions (4 nm). What is less clear is how PS I might be excluded from the grana partitions; its total mass is not terribly different from that of PS 11, although the aqueous projection may be. Perhaps it is highly negatively charged. How smaller extrinsic proteins like the ferredoxin-NADP reductase are excluded from grana partitions is also an open question. In summary, during the past few years the chloroplast has yielded much insight into the connection between protein mobility and membrane differentiation, but central pieces of the puzzle are still missing. In particular, the mechanism for lateral exclusion of PS I remains to be determined. In vitro stacking and

PROTEIN DIFFUSION IN CELL MEMBRANES

71

destacking may provide fertile ground for studying the role of lateral diffusion in membrane morphogenesis. 3. Phagocytosis There is an apparent sorting problem involved in leukocyte phagocytosis that passive redistribution models do not help to explain. While Fc, receptor activity is selectively lost from the murine macrophage cell surface during phagocytosis (Schmidt and Douglas, 1972; Petty et af., 1980a), other membrane proteins are retained, e.g., the receptor for complement component C3b, certain lectin receptors, and transport sites for lysine and adenosine (Tsan and Berlin, 1971; Ukena and Berlin, 1972; Oliver et al., 1974; Petty et al., 1980a). Although cell surface retention of a particular protein may be achieved through mechanisms as, for example, uptake followed by reinsertion of the same species from intracellular membranes or the unmasking of latent activities in the remaining membrane, it has been suggested that in some cases active cytoskeleton-mediated exclusion of selected proteins from the phagocytic membrane is involved (summary in Oliver and Berlin, 1982). Tsan and Berlin (1971) showed that irreversible inactivation of transport sites with a nonpenetrating reagent led to loss of transport activity (which was not recovered after phagocytosis), apparently indicating that new sites are not reinserted from inside the cell. Microtubule disrupting drugs prevent retention of adenine and lysine transport activities during phagocytosis (Ukena and Berlin, 1972), thus implicating some form of microtubule intervention. In a result bearing striking resemblance to that of Flanagan and Koch (1978) that cross-linked surface Ig binds to actin in lymphocytes, Jack and Fearon (1983) have now discovered that cross-linkage of C3b receptors in human PMNs leads to their association with a detergent (NP 40)-insoluble cytoskeleton fraction. Independent support for cytoskeletal control comes from the observation that initially randomly dispersed C3b receptors cluster and the clusters undergo oriented motion upon contact of polymorphonuclear leukocytes with various artificial substrata (Hafeman et a l . , 1982). Like anchorage modulation this is another global effect in response to local contact; clusters form over the entire surface upon attachment of one side of the cell to the substrate. Several lines of evidence thus substantiate the notion that active processes involving the cytoskeleton coordinate the exclusion of specific proteins from the phagocytic membrane.

IV. Closing Remarks Is diffusion of membrane proteins crucial for any known biological processes? The answer cannot be a resounding “yes” for all the membrane-directed events examined in this article. The problem stems from (1) a considerable uncertainty regarding the true diffusion rates of many proteins in cell membranes; (2) a lack

72

MICHAEL MCCLOSKEY AND MU-MING PO0

of more concrete knowledge of the context within which the proteins are moving. Without a sharper picture of membrane microstructure, even the most accurate of diffusion measurements is difficult to interpret. Quantitative analysis of many problems involving “short-range’’ motion of proteins is particularly handicapped by the spatial resolution of currently available tools. We have considered how lateral diffusion of membrane proteins may directly facilitate a number of membrane-mediated biological functions, including selfassembly, cell-cell recognition and adhesion, enzymatic reactions, hormonal response, and other signaling processes. Yet it is obvious that motional phenomena such as lateral diffusion do not owe their existence to the fact that they might subserve various membrane functions; on the contrary, the diffusion of a membrane protein must to some extent be a mere reflection of the fluid lipid environment in which it resides. In fact, unbridled protein diffusion represents a potentially disruptive force which the cell must contend with in structuring membrane topography during growth, differentiation, and normal functioning. We have seen through our discussion of diffusion-mediated trapping that mechanisms may have evolved which can channel this potentially negative force and make it work for the cell. It is our feeling that this economical device is perhaps the most intriguing aspect of protein diffusion in cell membranes. REFERENCES Abercrombie, M., Heaysman, J. E. M., and Pegrum, S. M. (1970). Exp. Cell Res. 62, 389-398. Adam, G., and Delbruck, M. (1968). In “Structural Chemistry and Molecular Biology” (A. Rich and N. Davidson, eds.), pp. 198-215. Freeman, San Francisco, California. Ahmed, A. J., Smith, H. T., Smith, M. B., and Millet, F. S. (1978). Biochemistry 17,2479-2483. Aksiyote-Benbasat, J., and Bloomfield, V. A. (1981). Biochemistry 20, 5018-5025. Albertsson. P. A., Hsu, B. D., Tang, G. M-s., and Amon, D. I. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 3971-3975. Albrecht-Buehler, G . (1981). Cold Spring Harbor Symp. Quant. Biol. 46, 45-49. Anderson, J . M. (1975). Biochim. Biophys. Acta 416, 191-235. Anderson, J . M. (1982). FEES Lett. 138, 62-66. Anderson, J . M., and Anderson, B. (1982). Trends Biochem. Sci. 7 , 288-292. Anderson, J . M., and Melis, A. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 745-749. Anderson, J. M., Goodchild, D. G., and Boardman, N. K. (1973). Biochim. Biophys. Acta 325, 573-585. Anderson, M.J., and Cohen, M. W. (1977). J. Physiol. (London) 268, 757-773. Anderson, M. J . , and Famrough, D. M. (1982). J. Cell B i d . 95, 120a. Argyroudi-Akoyunoglou, J. H., and Akoyunoglou, G. (1977). Arch. Biochem. Eiophys. 179, 370-377. Armond, P. A,, Amtzen, C. J., Briantais, J.-M., and Vemotte, C. (1976). Arch. Biochem. Eiophys. 175, 54-63. Amon, D. I., Tsujimoto, H. Y.,and Tang, G. M . 4 . (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 2942-2946. Amon, D. I., Hsu,B. D., Tang, G. M.-s., and Albertsson, P.-A. (1983). Fed. Proc. Fed. Am. Soc. Exp. Biol. 42, 2175.

PROTEIN DIFFUSION IN CELL MEMBRANES

13

Arntzen, C. J., Armond, P. A., Briantis, J.-M., Burke, J. J., and Novitsky, W. P. (1976). Brookhuven Symp. Biol. 28, 316-337. Avivi, A., Tramontano, D., Ambesi-Impiombato, F. S., and Sclessinger, J. (1981). Science 214, 1237-1239. Axelrod, D. (1983). J. Membr. Biol. 75, 1-10. Axelrod, D., Radvin, R., Koppel, D. E., Schlessinger, J., Webb, W. W., Elson, E. L.,and Podleski (1976). Proc. Nurl. Acud. Sci. U.S.A. 73, 4594-4598. Axelrod, D., Wight, A., Webb, W., and Honvitz, A. (1978). Biochemistry 17, 3604-3609. Azzi, A., Bill, K., and Broger, C. (1982). Proc. Nurl. Acad. Sci. U.S.A. 79, 2447-2450. Balakrishnan, K., Hsu, F. J., Cooper, A. D., and McConnell, H. M. (1982). J . Biol. Chem. 257, 6427-6433. Baldwin, D., Tenis, S., and Steiner, D. F. (1980). J. Biol. Chem. 255, 4028-4034. Barak, L. S., and Webb, W. W. (1981). Biophys. J. 33, 74a. Barak, L. S., and Webb, W. (1982). J . Cell Biol. 95, 846-853. Barber, J. (1980). FEES Len. 118, 1-10, Barisas, B. G., and Leuther, M. D. (1979). Biophys. Chem. 10, 221-229. Bayne, E. K., Anderson, M. J., and Fambrough, D. M. (1982). J . Cell Biol. 95, 117a. Bell, G. I. (1979). In “Physical Chemical Aspects of Cell Surface Events in Cellular Regulation,” pp. 371-385. Elsevier, Amsterdam. Bennet, J., Steinback, K. E., and Arntzen, C. J. (1980). Proc. Nurl. Acud. Sci. U.S.A. 77, 5255-5257. Bennett, V., and Stenbuck, P. J. (1979). J. Biol. Chem. 254, 2533-2541. Bennoun, P., and Jupin, H. (1974). Proc. Inr. Congr. Phorosynth., 3rd I, 163-169. Berclaz, T., and McConnell, H. M. (1981). Biochernisrry 20, 6635-6640. Berg, H. C., and Purcell, E. M. (1977). Biophys. J. 20, 193-219. Bergmann, J. E., Kupfer, A., and Singer, S. J. (1983). Proc. Nurl. Acad. Sci. U.S.A. 80, 1367-1371. Berzborn, R. J. (1969). Z. Nururforsch. UB, 436-446. Biggins, J. (1982). Biochim. Biophys. Acru 679, 479-482. Birdwell, C. R., Strauss, E. G., and Strauss, J. H. (1973). Virology 56, 429-438. Blackshaw, S., and Wamer, A. (1976). Nature (London) 262, 217-219. Bonaventura, C., and Myers, J. (1969). Biochim. Biophys. Actu 189, 366-383. Brasitus, T. A. (1983). Biochim. Biophys. Actu 728, 20-30. Bretscher, M. S. (1976). Nature (London) 260, 21-23. Bretscher, M. S., Thomson, J. N., and Pearse, B. M. F. (1980). Proc. Narl. Acud. Sci. U.S.A. 77, 4156-41 59. Brown, M. C., Jansen, J. K., and Van Essen, D. (1976). J. Physiol. 261, 387-422. Brown, M. F., Milajanich, G. P., and Dratz, E. A. (1977). Biochemistry 12, 2640-2648. Burke, J. J., Steinback, K. E., and Arntzen, C. J. (1979). Planr Physiol. 63, 237-244. Butler, W. L. (1978). Annu. Rev. Planr Physiol. 29, 345-378. Cadman, A. D., Fleming, R., and Guy, R. H. (1981). Biophys. J. 37, 569-574. Cameron, D. G., Martin, A., and Mantsch, H. H. (1983). Science 219, 180-182. Capaldi, R. A. (1982). Biochim. Biophys. Actu 694, 291-306. Caplan, A. I . , and Greenwalt, J. W. (1966). J . Cell. Biol. 31, 455-472. Cappelletti, R., Del Rosso, M., Vannucchi. S., Cella, C., Fibbi, G., and Chiarugi, V. P. (1980). J . Reticuloendothel. SOC. 27, 383-391. Carpentier, I.-L., Gordon, P., Anderson, R. G. W., Goldstein, J. L., Brown, M. S., Cohen, S., and Orci, L. (1982). J. Cell Biol. 95, 73-77. Cartwright, B., Smale, C. I.. Brown, F., and Hull, R. (1972). J. Virol. 10, 256-260. Cartwright, G. S., Smith, L. S . , Heinzelmann, E. W., Ruebush, M. J., Parce, J. W., and McConnell, H. M. (1982). Proc. Nurl. Acud. Sci. U.S.A. 79, 1506-1510.

74

MICHAEL MCCLOSKEY AND MU-MING P O 0

Chabay, R., DeLisi, C., Hook, W. A., and Siraganian, R. P. (1980). J . B i d . Chem. 255, 4628-4635. Chance, B., and Williams, G. R. (1956). Adv. Enzymol. 17, 65-130. Chang, C-h., Takeuchi, H., Ito, T., Machida, K., and Ohnishi, S. (1981). J. Biochem. (Tokyo)90, 997-1004. Chao, N-m., Young, S. H., and Poo, M.-m. (1981). Biophys. J. 36, 139-153. Chazotte, B., Wu, E.-s., and Hackenbrock, C. R. (1983). Fed. Proc. Fed. Am. SOC. Exp. B i d . 42, 2170. Cereijido, M., Ehrenfeld, J., Meza, I., and Martinez-Palomo, A. (1980). J. Membr. Biol. 52, 147- 159. Cherry, R. J. (1979). Biochim. Biophys. Actu 559, 289-327. Cherry, R. J., Busslinger, M., Schneider, G., and Parish, G. R. (1976). Narure (London) 263, 389-393. Cherry, R. J., Muller, U., and Schneider, G. (1977). FEES Leu. 80, 465-469. Cherry, R. J . , Muller, U., Holenstein, C., and Heyn, M. P. (1980). Biochim. Biophys. Acra 596, 145- 15 I . Chiarugi, V. P., and Vannucchi, S. (1976). J. Theor. Biol. 61, 459-475. Chothia, C., and Janin, J. (1975). Nature (London) 256, 705-708. Chow, I. (1980). Doctoral thesis, McGill University, Montreal. Chow, I., and Poo, M.-m. (1982). J . Cell Biol. 95, 510-518. Cohen, M. H., and Turnbull, D. (1959). J. Chem. Phys. 31, 1164-1 169. Cohen, M. W., and Weldon, P. R. (1980). J . Cell Biol. 86, 338-346. Cohen, M. W., Anderson, M. J., Zorychta, E., and Weldon, P. R. (1979). Prog. Brain Res. 49, 335-349. Cox, R. P., and Anderson, B. (1981). Biochem. Biophys. Res. Commun. 103, 1336-1342. Cutler, L. S. (1982). J . Cell Biol. 95, 130a. Davis, D. J., Armond, P. A , , Gross, E. L., and Arntzen, C. J . (1976). Arch. Biochem. Biophys. 175, 64-70. DeLisi, C. (1979). In “Physical Chemical Aspects of Cell Surface Events in Cellular Regulation,” pp. 261-292. Elsevier, Amsterdam. DeLisi, C. (1981). Mol. Immunol. 18, 507-511. DeLisi, C., and Wiegel, F. W. (1981). Proc. Narl. Acad. Sci. U.S.A. 78, 5569-5572. Dembo, M., Goldstein, B., Sobotka, A. K., and Lichtenstein, L. M. (1979a). J. Irnmunol. 122, 5 18-528. Dembo, M., Goldstein, B., Sobotka, A. K., and Lichtenstein, L. M. (1979b). J. Immunol. 123, 1864- 1872. de Petris, S . (1977). In “Dynamic Aspects of Cell Surface Organization” ( G . Poste and G. L. Nicolson, eds.), pp. 643-728. Elsevier, Amsterdam. Derzko, Z., and Jacobson, K. (1978). Biophys. J . 21, 204a. Deutsch, V. (1976). Virology 69, 607-616. Diamant, B., Kruger, P. G., and Uvnas, B. (1970). Acfu. Physiol. Scand. 79, 1-5. Dickson, R. B., Willingham, M. C., and Pastan, I. (1981). J. CellBiol. 89, 29-34. Dintzis, H. M., Dintzis, R. Z., and Vogelstein, B. (1976). Proc. Nafl. Acad. Sci. U.S.A. 73, 367 I -3675. Dintzis, R. Z., Vogelstein, B., and Dintzis, H. M. (1982). Proc. Natl. Acad. Sci. U.S.A. 79, 884-888. Dixit, B. P., Sudha, N., Waring, A. J., Wells, K. O., Wong, P. S., Woodrow, G. V., and Vanderkooi, J. M . (1982). Eur. J . Biochem. 126, 1-9. Dragsten, P. R., Blumenthal, R., and Handler, J. S. (1981). Nafure (London) 294, 718-722. Edelman, G. M. (1976). Science 192, 218-226.

PROTEIN DIFFUSION IN CELL MEMBRANES

75

Edidin, M. (1981). In “Membrane Structure’’ (J. B. Finean and R. H. Michell, eds.), pp. 37-82. Elsevier, Amsterdam. Edidin, M., and Fambrough, D. (1973). J. Cell Biol. 57, 27-37. Edidin, M., and Wei, T. (1982). J. Cell Biol. 95, 458-462. Edidin, M., Wei, T., and Gotlib, L. (1982). J . Cell Biol. 95, 453-457. Edwards, C., and Frisch, H. L. (1976). J . Neurobiol. 7, 377-381. Elgsaeter, A., Shotton, D. M.. and Branton, D. (1976). Biochim. Biophys. Acta 426, 101-122. Emeis, C. A., and Fehder, P. L. (1970). J. Am. Chem. Soc. 92, 2246-2252. Erecinska, M., Davis, J. S., and Wilson, D. F. (1980). J. Biol. Chem. 255, 9653-9658. Ernster, L., Carlsson, C., Hundal, T., and Nordenbrand, K. (1979). Methods Enzymol. 55, 399-407. Evans, E., and Buxmbaum, K. (1981). Biophys. J. 34, 1-12. Evans, E. A,, and Hochmuth, R. M. (1978). Curr. Top. Membr. Tramp. 10, 1-64. Evans, W. (1980). Biochim. Biophys. Acta 604, 27-64. Fahey, K. R., and Green, A. A. (1938). J. Am. Chem. SOC. 60, 3039-4043. Ferguson-Miller, S., Brautigan, D. L., and Margoliash, E. (1978). J. Biol. Chem. 253, 149-159. Flanagan, J., and Koch, G. L. E. (1978). Nature (London) 273, 278-281. Fowler, V., and Branton, D. (1977). Nature (London) 268, 23-26. Fraser, S . E., and Po0 M-m. (1982). Curr. Top. Dev. Biol. 17, 77-100. Frye, L. D., and Edidin, M.(1970). J. Cell Sci. 7, 319-335. Galla, H.-J., and Sackmann, E. (1974). Biochim. Biophys. Actu 339, 103-1 15. Galli, P., Brenna, A., De Camilli, P., and Meldolesi, J . (1976). Exp. Cell Res. 99, 178-183. Gershon, N. D. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 1357-1360. Gingell, D. (1976). In “Mammalian Cell Membranes” (G. A. Jamieson and D. M. Robinson, eds.), pp. 198-223. Butterworths, London. Gingell, D., and Ginsberg, L. (1978). In “Membrane Fusion” (G. Poste and G. Nicolson, eds.), pp. 792-833. Elsevier, Amsterdam. Golan, D. E., and Veatch, W. (1980). Proc. Nurl. Acud. Sci. U . S . A . 77, 2537-2541. Golan, D. E., and Veatch, W. R. (1981). Report at the International Workshop on the Biological Applications of Photobleaching Techniques, Chapel Hill, North Carolina. Goldstein, B., Wofsy, C . , and Bell, G. (1981). Proc. Nut/. Acud. Sci. U.S.A. 78, 5695-5698. Goldstein, J. L., Anderson, R. G. W., and Brown, M. S. (1979). Nature (London) 279, 679-685. Goodenough, U. W., and Staehelin, L. A. (1971). J. Cell. Biol. 48, 594-619. Gottlieb, C., Kornfeld, S., and Schlesinger, S. (1979). J. Virol. 29, 344-351. Gruler, H. (1975). Z. Narurforsch. MC, 608-614. Gupte, S., Jacobson, K., Hoechli, L., and Hackenbrock, C. R. (1983). Biophys. J . 41, 371a. Hackenbrock, C. R. (1981). Trends Biochem. Sci. 6, 151-154. Hackenbrock, C. R., and Hammon, K. M. (1975). J. Biol. Chem. 250, 9185-9197. Haehnel, W., Hesse, V., and Propper, A. (1980). FEES Lett. 111, 79-82. Hafeman, D. G., von Tscharner, V., and McConnell, H. M. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 4552-4556. Hafeman, D. G., Smith, L. M., Fearon, D. T., and McConnell, H. M. (1982). J. Cell Biol. 94, 224-227. Haines, T. H. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 160-164. Haley, B., Hoyer, P. B., and Middaugh, C. R. (1983). Fed. Proc. Fed. Am. Soc. Exp. Biol. 42, 1851. Hanski, E., Rimon, G . , and Levitzki, A. (1979). Biochemistry 18, 846-853. Hardt, S. L. (1979). Biophys. Chem. 10, 239-243. Hams, A. K. (1976). Nature (London) 263, 781-783. Hart, G. W. (1982). Biochemistry 21, 6088-6096.

76

MICHAEL MCCLOSKEY AND MU-MING PO0

Hatefi, Y. (1968). Proc. Narl. Acad. Sci. U.S.A. 60, 733-740. Haworth, P., Kyle, D. J., Horton, P., and Arntzen, C. J. (1982). Phorochem. Phorobiol. 36, 743-748. Haywood, A. M. (1975). J. Gen. Virol. 29, 63-68. Hazum, E., Chang, K.-j., and Cuatrecasas, P. (1980). Proc. Narl. Acad. Sci. U.S.A. 77, 3038-3041. Heath, J . P. (1983). Narure (London) 302, 532-534. Henis, Y. I., and Elson, E. L. (1981). Proc. Narl. Acad. Sci. U.S.A. 78, 1072-1076. Henis, Y. I., and Elson, E. L. (1983). Personal communication. Henis, Y.I., Hekman, M., Elson, E. L., and Helmreich, E. J. M. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 2907-291 1. Hillman, G. M., and Schlessinger, J. (1982). Biochemistry 21, 1667-1672. Hirata, F., and Axelrod, J . (1980). Science 209, 1082-1090. Hochman, J. H., Schindler, M., Lee, J. G., and Ferguson-Miller, S. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 6866-6870. Hochman, J., Ferguson-Miller, S., Foxall, S., and Schindler, M. (1983). Eiophys. J. 41, 6a. Horton, P., and Black, M. T. (1980). FEES Leu. 119, 141-144. Horton, P., and Croze, E. (1979). Eiochim. Eiophys. Acra 545, 188-201. Hubbell, W. L., and McConnell, H. M. (1968). Proc. Narl. Acad. Sci. U.S.A. 61, 12-16. Hubbell, W. L., and McConnell, H. M. (1969). Proc. Narl. Acad. Sci. U.S.A. 64, 20-27. Hughes, B. D., Pailthorpe, B. A., White, L. R., and Sawyer, W. H. (1982). Biophys. J . 37, 673-676. Ingram, V. M. (1969). Narure (London) 222, 641-644. Insel, P. A., and Kennedy, M. S. (1978). Narure (London) 273, 471-473. Isersky, C., Taurog, J., Poy, G., and Metzger, H. (1978). J . Immunol. 121, 549-558. Ishizaka, T. (1982). Fed. Proc. Fed. Am. SOC. Exp. Eiol. 41, 17-21. Ishizaka, T., and Ishizaka, K. (1978). J . Immunol. 120, 800-805. Ishizaka, T., Tomioka, H., and Ishizaka, K. (1971). J . Immunol. 106, 705-710. Izawa, S., and Good, N. (1966). Plant Physiol. 41, 544-552. Jack, R. M., and Fearon, D. T. (1983). Fed. Proc. Fed. Am. SOC. Exp. Eiol. 42, 1235. Jacobson, K. (1980). In “Lasers in Biology and Medicine” (F. Hillenkamp, R. Pratesi, and C. A. Sacchi, eds.), pp. 271-288. Plenum, New York. Jacobson, K., and Wojcieszyn, J . (1981). Comments Mol. Cell. Eiophys. 1, 189-199. Jacobson, K., Hou, Y., Derzko, Z., Wojcieszyn, J., and Organisciak, D. (1981). Eiochemisrry 20, 5268-5275. Jennings, R. C., Garlaschi, F. M., Gerola, P. D., and Forti, G . (1979). Eiochim. Biophys. Acru 456, 207-2 19. Jesaitis, A. J., and Yguerabide, J. (1981). Report at the “International Workshop on the Biological Applications of Photobleaching Techniques,” Chapel Hill, North Carolina. Johnson, D. C., and Schlesinger, M. J. (1980). Virology 103, 407-424. Johnson, D. C., Schlesinger, M. J., and Elson, E. L. (1981). Cell 23, 423-431. Johnson, P., and Garland, P. B. (1981). FEBS Lett. 132, 252-256. Kagey-Sobotka, A., MacGlashan, D. W., and Lichtenstein, L. M. (1982). Fed. Proc. Fed. Am. SOC. Exp. Biol. 41, 12-16. Kahn, C. R. (1979). Proc. SOC. Exp. Eiol. Med. 162, 13-21. Kampe, O., and Peterson, P. A. (1979). Report at the Soviet-Swedish Symposium on Chemistry and Physics of Biological Membranes, Stockholm-Lidingo. Kawato, S., Sigel, E., Carafoli, E., and Cheny, R. J. (1980). J . Biol. Chem. 255, 5508-5510. Kawato, S., Sigel, E., Carafoli, E., and Cherry, R. J. (1981). J. Eiol. Chem. 256, 7518-7527.

PROTEIN DIFFUSION IN CELL MEMBRANES

77

Ketis, N. V., and Grant, C. W. M. (1982). Biochim. Biophys. Acfu 685, 347-354. Ketis, N. V., Girdlestone, J., and Grant, C. W. M. (1980). Proc. Nurl. Acud. Sci. U.S.A. 77, 3788-3790. Kimelberg, H. K. (1977). In “Dynamic Aspects of Cell Surface Organization” (G. Poste and G. L. Nicolson, eds.), pp. 205-293. Elsevier, Amsterdam. Kjellen, I., Pettersson, I., and Hook, M. (1981). Proc. Nurl. Acud. Sci. U.S.A. 78, 5371-5375. Klausner, R. D., and Wolf, D. E. (1980). Biochemistry 19, 6199-6203. Klausner, R. D., Bhalla, D. K., Dragsten, P., Hoover, R. L., and Kamovsky, M. J. (1980). Proc. Nufl. Acud. Sci. U.S.A. 77, 437-441. Klingenberg, M. (1967). In “Mitochondria1 Structure and Compartmentation” (E. Quagliariello, S . Papa, E. C. Slater, and J. M. Tager, eds.), pp. 124-125. Adriatica Editrice, Bari, Italy. Knipe, D. M., Baltimore, D., and Lodish, H. F. (1977). J. Virol. 21, 1128-1139. Kojima, K., and Koizumi, K. (1974). In?. Cancer Congr., IIrh, Florence Oct. 21, 1974. Koppel, D. E . , Sheetz, M. P., and Schindler, M. (1981). Proc. Nurl. Acud. Sci. U.S.A. 78, 3576-3580. Koppel, D. E., Oliver, J. M., and Berlin, R. D. (1982). J . Cell Biol. 93, 950-960. Kraemer, P. M., and Tobey, R. A. (1972). J . Cell Biol. 55, 713-717. Kroger, A., and Klingenberg, M. (1973a). Eur. J . Biochem. 34, 358-368. Kroger, A., and Klingenberg, M. (1973b). Eur. J . Biochem. 39, 313-323. Kyle, D. J . , Staehelin, L. A., and Amtzen, C. J. (1983). Arch. Biochem. Biophys. 222,527-541. Laggner, P. (1981). Nurure (London) 294, 373-374. Lawson, D., Fewtrell, C., and Raff, M. C. (1978). J . Cell Biol. 79, 394-400. Lazarowitz, S. G.,Compans, R. W., and Choppin, P. W. (1973). Virology 52, 199-212. Lee, P. M., and Grant, C. W. M. (1979). Biochem. Biophys. Res. Commun. 90, 856-863. Lehninger, A. L. (1959). Rev. Mod. Phys. 31, 136-146. Lenard, J. (1978). Annu. Rev. Biophys. Bioeng. 7, 139-165. Levine, J . , and Willard, M. (1983). Proc. Nutl. Acud. Sci. U.S.A. 80, 191-195. Lewis, J. T., Hafeman, D. G.,and McConnell, H. M. (1980). Biochemistry 19, 5376-5386. Lichtman, M. A. (1970). N . Engl. J . Med. 283, 943-948. Liebman, P. A,, and Entine, G.(1974). Science 185, 457-459. Lin, F. H., and Thormar, H. (1980). Nature (London) 285, 490-492. Little, S . P., and Huang, A. S . (1977). Virology 81, 37-47. Luft, J. H. (1976). Inr. Rev. Cyrol. 45, 291-382. Luna, E. J . , Kidd, G.H., and Branton, D. (1979). J . Biol. Chem. 254, 2526-2532. Malveaux, F. J . , Conroy, M. C., Adkinson, N. F., and Lichtenstein, L. M. (1978). J . Clin. Invest. 62, 176-181. Marcelja, S. (1976). Biochim. Biophys. Actu 455, 1-7. Marsh, D., Watts, A,, Pates, R. D., Uhl, R., Knowles, P. F., and Esmann, M. (1982).Biophys. J . 37, 265-274. Matsuura, S . , Nakada, H., Sawamura, T., and Tashiro, Y. (1982). J . Cell Biol. 95, 864-875. Maxfield, F. R., Schlessinger, J., Schecter, Y., Pastan, I., and Willingham, M. C. (1978). Cell 14, 805-8 10. McConnell, H. M. (1979). In “Membranes and Intercellular Communication” (R. Balian, M. Chabre, and P. F. Devaux, eds.), pp. 270-292. North Holland Publ., Amsterdam. Melis, A,, and Hohman, P. H. (1976). Phorochem. Phorobiol. 23, 343-350. Mellman, I . , Unkeless, J., Steinman, R., and Cohn, Z. (1981). J. Cell Biol. 91, 124a. Mendoza, G.,and Metzger, H. (1976). Nurure (London) 264, 548-550. Metcalf, T. N., Schindler, M., Schubert, K. R., and Wang, J. L. (1982). J . Cell Biol. 95, 463a. Michell, R . H., Coleman, R., and Lewis, B. A. (1976). Biochem. SOC. Trans. 4, 1017-1020.

78

MICHAEL MCCLOSKEY AND MU-MING PO0

Michl, J . , Unkeless, J., Pieczonka, M. M., and Silverstein, S. C. (1979). J. Cell Eiol. 83, 295a. Michl, J . , Pieczonka. M. M., Unkeless, J., Bell, G., and Silverstein, S . C. (1983). J. Exp. Med. 157, 2121-2139. Moreadith, R. W., and Jacobus, W. E. (1982). J. Biol. Chem. 257, 899-905. Mosbach, K . (1976). FEES Lett. 62 (Suppl.), E8O-E95. Muller, W. A,, Steinman, R. M., and Cohn, Z. A. (1980). J. Cell Eiol. 86, 304-314. Mullet, J . E., and Amtzen, C. J. (1980). Eiochim. Eiophys. Acta 589, 100-117. Murata, N. (1969). Eiochim. Eiophys. Acta 172, 242-251. Naqvi, K. R. (1974). Chem. Phys. Lett. 28, 280-284. Nicholls, P. (1976). Eiochim. Biophys. Acta 430, 30-45. Niedel, J . E., Kahane, I., and Cuatrecasas, P. (1979). Science 205, 1412-1414. Nigg, E. A,, and Cherry, R. J . (1980). Proc. Nut/. Acad. Sci. U.S.A. 77, 4702-4706. Ohyashiki, T., Takeuchi, M., Kodera, M., and Mohri, T. (1982). Biochim. Biophys. Acta 688, 16-22. Ojakian, G. K., and Satir, P. (1974). Proc. Natl. Acad. Sci. U.S.A. 71, 2052-2056. Olden, K . , Parent, J . B., and White, S . L. (1982). Biochim. Biophys. Acta 650, 209-232. Oliver, J . M., and Berlin, R. D. (1982). Int. Rev. Cytol. 74, 55-94. Oliver, J . M., Ukena, T. E., and Berlin, R. D. (1974).Proc. Nail. Acad. Sci. U.S.A. 71, 394-398. Olmsted, J. B., and Borisy, G. G. (1973). Annu. Rev. Eiochern. 42, 507-540. Orci, L., Carpentier, J.-L., Perrelet, A., Anderson, R. G. W., Goldstein, 1. L., and Brown, M. S . (1978). Exp. Cell Res. 113, 1-13. Orly, J., and Schramm, M. (1976). Proc. Nail. Acad. Sci. U.S.A. 73, 4410-4414. Overfield, R. E., and Waight, C. A. (1980). Biochemistry 19, 3328-3334. Owicki, J . C., and McConnell, (1979). Proc. Nail. Acad. Sci. U.S.A. 76, 4750-4754. Owicki, J. C., and McConnell, H. M. (1980). Eiophys. J . 30, 383-397. Owicki, I. C., Springgate, M. W., and McConnell, H. M. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 16 16- 16 19. Parr, E. L., and Kirby, W. N. (1979). J. Histochem. Cytochem. 27, 1327-1336. Peters, R. (1981). Cell Eiol. Int. Rep. 5, 733-760. Peters, R., and Cherry, R. J. (1982). Eiophys. J . 37, 277a. Petersen, N. 0..McConnaughey, W. B., andElson, E. L. (1982). Proc. Nail. Acad. Sci. U.S.A. 79, 5327-533 I . Petit, V. A., and Edidin, M. (1974). Science 184, 1183-1185. Petty, H. R., Hafeman, D. G., and McConnell, H. M. (1980a). J. Immunol. 125, 2391-2396. Petty, H. R., Smith, L. M . , Fearon, D. T., and McConnell, H. M. (1980b). Proc. Nail. Acad. Sci. U.S.A. 77, 6587-6591. Petty, H. R., Hafeman, D. G., and McConnell, H. M. (1981). J . Cell Eiol. 89, 223-229. Pfeiffer, J. R., Oliver, J. M., and Berlin, R. D. (1980). Nature (London) 286, 727-729. Pinto, da Silva, P., and Nicolson, G. L. (1974). Eiochim. Eiophys. Acra 363, 311-319. Pisam, M., and Ripoche, P. (1976). J. Cell B i d . 71, 907-920. Podesta, E. J., Solano, A. R., Attar, R., Senchez, M. L., and Molina y Vedia, L. (1983). Proc. Nut/. Acad. Sci. U.S.A. 80, 3986-3990. Poo, M-m. (1981). Annu. Rev. Eiophys. Bioeng. 10, 245-276. Poo, M-m. (1982). Nature (London) 295, 332-334. Poo, M-m., and Cone, R. A. (1974). Nature (London) 247, 438-441. Poo, M-m.,Lam, J . W., Orida, N., and Chao, A. W. (1979). Biophys. J . 26, 1-21. Popp, C. A,, and Hyde, J. S . (1982). Proc. Nail. Acad. Sci. U.S.A. 79, 2559-2563. Poulos, T. L., and Kraut, J. (1980). J. Biol. Chem. 255, 10322-10330. Punnet, T. (1971). Science 171, 284-286. Purves, D., and Lichtman, J . W. (1980). Science 210, 153-157.

PROTEIN DIFFUSION IN CELL MEMBRANES

79

Quinn, P. 1. (1981). f r o g . Biophys. Mol. Biol. 38, 1-104. Rambourg, A. (1971). Int. Rev. Cytol. 31, 57-114. Rasenick, M. M., Stein, P. J . , and Bitensky, M. W. (1981). Nature (London) 294, 560-562. Razi-Naqvi, K., Behr, J.-P., and Chapman, D. (1974). Chem. Phys. Leu. 26, 440-444. Reidler, J. A,, Keller, P. M., Elson, E. L., and Lenard, J . (1981). Biochemistry 20, 1345-1349. Reger, B. J., and Kraus, R. W. (1970). Plant Physiol. 46, 568-575. Revesz, T., and Greaves, M. (1975). Nature (London) 257, 103-106. Richter, P. H., and Eigen, M. (1974). Biophys. Chem. 2, 255-263. Rieder, R., and Bosshard, H. R. (1980). J . B i d . Chem. 255, 4732-4739. Rimle, D., Morse, P. D., and Njus, D. (1983). Biochim. Biophys. Acta 728, 92-96. Rizki, T. M., and Rizki, R. M. (1983). Nature (London) 303, 340-342. Rodbell, M. (1980). Nature (London) 284, 17-22. Rodbell, M. (1981). In “Membranes and Intercellular Communication” (R. Balian, M. Chabre, and P. F. Devaux, eds.), pp. 369-385. North Holland Publ., Amsterdam. Roth, T. F., and Woods, J. W. (1982). “Differentiation and Function of Hematopoietic Cell Surfaces,” pp. 163-181. Liss, New York. Rothmann, J. E. (1981). Science 213, 1212-1220. Rothmann, J. E., Fries, E., Dunphy, W. G., and Urbani, L. J. (1981). Cold Spring Harbor Symp. Quant. Biol. 46, 797-805. Rubenstein, J. L. R., Owicki, J. C., and McConnell, H. M. (1980). Biochemistry 19, 569-573. Rubin, B. T., Barber, J., Paillotin, G., Chow, W. S., and Yamamoto, Y. (1981). Biochim. Biophys. Acta 638, 69-74. Rudolph, S. A,, Hegstrand, L. F., Greengard, P., and Malawista, S. E. (1979). Mol. Pharmacol. 16, 805-812. Saffman, P. G., and Delbruck, M. (1975). Proc. Natl. Acad. Sci. U.S.A. 72, 31 11-31 13. Sampaio, L. 0.. Dietrich, C. P., and Giannotti, F. (1977). Biochim. Biophys. Acta 498, 123-131. Sanes, J. R., Marshall, L. M., and McMahan, U. J. (1978). J . Cell B i d . 78, 176-198. Sang, U. H., Saier, M. H., and Ellisman, M. H. (1979). Exp. Cell Res. 122, 384-391. Saxton, M. J. (1982). Biophys. J. 39, 165-173. Schechter, Y., Chang, K. J., Jacobs, S., and Cuatrecasas, P. (1979a). Proc. Natl. Acad. Sci. U.S.A. 76, 2720-2724. Schechter, Y., Hemez, L., Schlessinger, J., and Cuatrecasas, P. (1979b). Nature (London) 278, 835-838. Schindler, M., Koppel, D. E., and Sheetz, M. P. (1980a). Proc. Natl. Acad. Sci. U.S.A. 77, 1457- 1461. Schindler, M., Osborn, M. J., and Koppel, D. E. (1980b). Nature (London) 285, 261-263. Schlessinger, J. (1979). In “Physical Chemical Aspects of Cell Surface Events in Cellular Regulation,” pp. 89-1 18. Elsevier, Amsterdam. Schlessinger, J., and Elson, E. L. (1981). In “Membrane Receptors: Methods for Purification and Characterization” (S. Jacobs and P. Cuatrecasas, eds.), pp. 159-170. Chapman & Hall, London. Schlessinger, J., Webb, W. W . , Elson, E. L., and Metzger, H. (1976). Nature (London) 264, 550-552. Schlessinger, J., Shechter, Y., Willingham, M. C., and Pastan, I. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 2659-2663. Schmidt, M. E., and Douglas, S. D. (1972). J. Immunol. 109, 914-917. Schmitz, K. S., and Schurr, J. M. (1972). J. Phys. Chem. 76, 534-545. Schneider, H., Lemasters, J. J . , Hochli, M., and Hackenbrock, C. R. (1980). J. B i d . Chem. 255, 3748-3756. Schneider, H., Hochli, M., and Hackenbrock, C. R. (1982). J. Cell B i d . 94, 387-393. Schnitzer, T. J., and Lodish, H. F. (1979). J . Virol. 29, 443-447.

80

MICHAEL MCCLOSKEY AND MU-MING P O 0

Schranner, R., and Richter, P. H. (1978). Biophys. Chem. 8, 135-150. Schroeder, F. (1980). Eur. J . Biochem. 112, 293-307. Schroeder, H. (1977). J. Chem. Phys. 67, 1617-1619. Schroit, A. J., and Pagano, R. E. (1981). Ce/l23, 105-112. Schulster, D. (1979). Trans. Biochem. Soc. 7, 310-314. Sedlacek, H. H., Stark, J.. Seiler, F. R., Ziegler, W., and Wiegandt, H. (1976). FEES Lerr. 61, 272-276. Segal, D. M., Taurog, J., and Metzger, H. (1977). Proc. Narl. Acad. Sci. U.S.A. 74, 2993-2997. Seul, M., Weiss, R. M., and McConnell, H. M. (1983). Biophys. J . 41, 212a. Sharom, F. J., and Grant, C. W. M. (1977). Biochem. Biophys. Res. Commun. 74, 1039-1045. Sharom, F. J., and Grant, C. W. M. (1978). Biochim. Biophys. Acfa 507, 280-293. Sheetz, M. P., Febbroriello, P., and Koppel, D. E. (1982). Nature (London) 296, 91-93. Shinitzky, M., and Henkart, P. (1979). Inr. Rev. Cyrol. 60, 121-147. Shoup, D., and Szabo, A. (1982). Biophys. J. 40,33-39. Siegel, C. O . , Jordan, A. E., and Miller, K. R. (1981). J. Cell Biol. 91, 113-125. Simmons, E. L. (1975). 2. Phys. Chem. 96, 47-58. Singer, S. J. (1976). In “Surface Membrane Receptors” (R. A. Bradshaw, W. A. Frazier, R. C. Merrell, D. 1. Gottlieb, and R. A. Hogue-Angeletti, eds.), pp. 1-24. Plenum, New York. Smith, H. G.,Fager, R. S., and Litman, B. J. (1977). Biochemistry 16, 1399-1405. Smith, L., Davies, H. C., and Nava, M. E. (1979a). In “Cytochrome Oxidase” (T. E. King, Y. Orii, B. Chance, and K. Okunuki, eds.), pp. 293-304. Elsevier, Amsterdam. Smith, L. M., Parce, J. W., Smith, B. A,, and McConnell, H. M. (1979b). Proc. Narl. Acud. Sci. U.S.A. 76, 4177-4179. Smith, L. M., Smith, B. A., and McConnell, H. M. (1979~).Biochemistry 18, 2256-2259. Smith, L. M., Rubenstein, J. L. R., Parce, J. W., and McConnell, H. M. (1980).Biochemistry 19, 5907-591 1 . Smith, L. M., Petty, H. R., Parham, P., and McConnell, H. M. (1982). Proc. Narl. Acad. Sci. U.S.A. 79, 608-612. Smithson, K. W., Millar, D. B., Jacobs, L. R., and Gray, G. M. (1981). Science 214, 1241-1243. Solc, K., and Stockmayer, W. H. (1973). Int. J . Chem. Kiner. 5, 733-752. Sowers, A. E., and Hackenbrock, C. R. (1981). Proc. Nufl. Acud. Sci. U.S.A. 78, 6246-6250. Speck, S. H., Ferguson-Miller, S . , Osheroff, N., and Margoliash, E. (1979). Proc. Nurl. Acud. Sci. U.S.A. 76, 155-159. Srere, P. A. (1982). Trends Biochem. Sci. 7, 375-378. Staehelin, L. A. (1976). J . Cell B i d . 71, 136-158. Staehlelin, L. A., Carter, D. P., and McDonnel, A. (1980).In “Membrane-Membrane Interactions” (N.B. Gilula, ed.), pp. 179-193. Raven, New York. Staehelin, L. A., Kyle, D. J . , and Arntzen, C. J. (1982). Plunr Physiol. 695, 69. Stem, P. L., and Bretscher, M. S. (1979). J . Cell Biol. 82, 829-833. Stonehuerner, J., Williams, J. B., and Millet, F. (1979). Biochemistry 18, 5422-5427. Storey, B. T., and Lee, C.-p. (1981). Trends Biochem. Sci. 6, 166-170. Stuhmer, W., and Almers, W. (1982). Proc. Narl. Acud. Sci. U.S.A. 79, 946-950. Sugrue, S. P., and Hay, E. D. (1981). J . Cell B i d . 91, 45-54. Takeuchi, J., Sobue, M., Sato, E., Shamoto, M., Miura, K., and Nakagaki, S. (1976). Cancer Res. 36, 2133-2139. Tanaka, K.-i., and Ohnishi, S.4. (1976). Biochim. Biophys. Acru 426, 218-231. Tank, D. W., Wu, E.-s., and Webb, W. W. (1982). J. CellBiol. 92, 207-212. Tartakoff, A. M. (1980). Inr. Rev. Exp. Purhol. 22, 227-251. Thielen, A. P. G. M., and Van Gorkom, H. J. (1981). Biochim. Biophys. Acra 635, 111-120. Thompson, D. A., Suarez-Villafane, M., and Ferguson-Miller, S. (1982). Biophys. J . 37,285-293.

PROTEIN DIFFUSION IN CELL MEMBRANES

81

Tolkovsky, A. M., and Levitsky, A. (1978). Biochemistry 17, 3795-3810. 147-170. Torney, D. C., and McConnell, H.M. (1983). Proc. R. Soc. (London) -7, Treffers, H. P. (1940). J. Am. Chem. SOC. 62, 1405-1409. Tsan, M.-f., and Berlin, R. D. (1971). Proc. Natl. Acad. Sci. U.S.A. 134, 1016-1035. Ukena, T. E., and Berlin, R. D. (1972). J. Exp. Med. 136, 1-7. Usharani, P.,Rao, L. V. M., Tokuyasu, K. T., and Butler, W. L. (1983). Eiophys. J . 41, 319a. Vaz, W. L. C., Kapitza, H.G., Stumpel, J., Sackmann, E., and Jovin, T. M. (1981). Biochemistry 20, 1392-1396. Wagner, R., Carrillo, N., Junge, W., and Vallejos, R. H.(1982). Biochim. Eiophys. Acta 680, 3 17-330. Wang, R. T., and Meyers, J. (1974). Biochim. Biophys. Acfa 347, 134-140. Wank, S. A., DeLisi, C., and Metzger, H.(1983). Biochemistry 22, 954-959. Waring, A., Davis, J. S., Chance, B., and Erecinska, M. (1980). J . Biol. Chem. 255,6212-6218. Weatherbee, J. A. (1981). Int. Rev. Cyrol. Suppl. 12, 113-176. Weaver, D. L. (1983). Biophys. J . 41, 81-86. Webb, W. W., Barak, L. S . , Tank, D. W., and Wu, E.-s. (1982). Biochem. SOC. Symp. 46, 191-205. Weigel, P. H. (1980). J. Cell Biol. 87, 855-861. Welch, G. R., and Gaertner, F. H. (1975). Proc. Natl. Acad. Sci. U.S.A. 72, 4218-4222. Wikstrom, M. (1981). Trends Biochem. Sci. 6, 166-170. Williams, R. J . P. (1981). Trends Biochem. Sci. 6, X. Willingham, M. C., Pastan, I. H., Sahagian, G. G., Jourdian, G. W., and Neufeld, E. F. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 6967-6971. Willison, J . H. M., Grout, B. W. W., and Cocking, E. C. (1971). Bioenergetics 2, 371-382. Wilson, T., and Lenard, J. (1981). Biochemistry 20, 1349-1354. Wise, B. C., Guidotti, A., and Costa, E. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 886-890. Wohlrab, H. (1970). Biochemistry 9, 474-479. Wu, E. S . , and Yang, C. S. (1980). Fed. Proc. Fed. Am. SOC. Exp. Biol. 39, 1990. Wu, E.-s., Jacobson, K., Szoka, F., and Portis, A. (1978). Biochemistry 17, 5543-5550. Wu, E. S., Low, P. S., and Webb, W. W. (1981). Biophys. J . 33, 109a. Wu, E.-s., Tank, D. W., and Webb, W. W. (1982). Proc. Natl. Acad. Sci. U.S.A. 79,4962-4966. Young, S . H.,and Poo, M.-m. (1983). J . Neurosci. 3, 225-23 1. Yu, J., and Branton, D. (1976). Proc. Narl. Acad. Sci. U.S.A. 73, 3891-3895. Yu, J., and Goodman, S. R. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 2340-2344. Ziomeck, C. A., Schulman, S., and Edidin, M. (1980). J. Cell Biol. 86, 849-857.