molecular architecture and functioning of the outer

Biochimica et Biophysica A cta, 737 (1983) 51 - 115 51 Elsevier Biomedical Press

BBA 85241

MOLECULAR ARCHITECTURE AND FUNCTIONING OF THE OUTER MEMBRANE OF ESCHERICHIA C O L I AND OTHER GRAM-NEGATIVE BACTERIA

BEN LUGTENBERG a,, and LOEK VAN ALPHEN h

" Department of Molecular Cell Biology' and Institute for Molecular Biology', State University, Transitorium 3, Padualaan 8, 3584 CH

Utrecht and h Laboratorium voor de Gezondheidsleer, University of Amsterdam, Mauritskade 57, 1092 AD Amsterdam (The Netherlands)

(Received July 26th, 1982)

Contents

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 A. Scope of this review . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 B. Ecological considerations relevant to structure and functioning of the outer membrane of Enterobacteriaceae . . . . . . . . 53 C. General description of the cell envelope of Gram-negative bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53

II. Methods for the isolation of outer membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 A. E. coli and S. typhimurium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58

1. Isolation of peptidoglycan-less outer membranes after spheroplast formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 2. Isolation of outer membrane-peptidoglycan complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 3. Differential membrane solubilization using detergents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 4. Membrane separation based on charge differences of vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59

B. Other organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59

II1. Individual constituents of the outer membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 A. Composition of the outer membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 B. Phospholipid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 C. Lipopolysaccharide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 2, Methods of isolation and purification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 3. Chemical structure of lipopolysaccharides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 4. Effects of polymixin, EDTA and divalent cations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

D. Enterobacterial common antigen (ECA) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64 E. Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

1. Introductory remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64 2. Enzymes in the outer membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 3. Lipoproteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68 4. OmpA protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 5. The family of peptidoglycan-associated general diffusion pore proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71

a. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 b. Function of general pore proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 c. Purification and properties of general diffusion pore proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 d. Characteristics of individual general diffusion pore proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

* To whom correspondence should be addressed. Abbreviations: Abe, abequose; ECA, Enterobacterial Common Antigen; ESR, electron spin resonance; GIcN, glucosamine; GIcNAc, N-acetyl-D-glucosamine; Hep. t-glycero-D-manno

heptose; KDO, 2-keto-3-deoxy-octulosonic acid; LPS, lipopolysaccharide; NMR, nuclear magnetic resonance; PAL, peptidoglycan-associated lipoprotein; Rha, rhamnose; SDS, sodium dodecyl sulphate.

0304-4157/83/0000-0000/$03.00 © 1983 Elsevier Science Publishers

52

6,

i. O m p C prote in and O m p F protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

ii. PhoE protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

iii. S a l m o n e l l a pore proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

iv. Other general diffusion pore proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

Charac ter i s t ics of E . c o i l pore prote ins not ant igenical ly re la ted to the family of pep t idoglycan-assoc ia ted general d i f fus ion pore prote ins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

a. Bacter iophage T6 receptor prote in . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

b. Bacter iophage l ambda receptor prote in . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

c. Outer m e m b r a n e pro te in involved in the up take of v i t amin B12 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

d. Outer m e m b r a n e prote ins involved in the up take of ferric ions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

Character is t ics of E . c o i l outer m e m b r a n e prote ins wi thout ident i f ied funct ion . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

a. Protein a . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

b. Protein II1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

c. LPS b ind ing pro te in . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

d. Outer m e m b r a n e prote ins induced by sulphate l imi ta t ion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

e. Phage- and p lasmid-coded outer m e m b r a n e prote ins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82

IV. Molecular organiza t ion of the outer membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 A. In t roduct ion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82

B. Methods used for s tudying the local iza t ion of ind iv idua l outer m e m b r a n e cons t i tuents and their in terac t ions . . . . . . . . 84

1. Local iza t ion at the cell surface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84

2. Prote in-prote in and pro te in -pep t idog lycan nearest ne ighbour associa t ions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85

3 In terac t ions be tween indiv idual prote ins and LPS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

4. The l ipid mat r ix and in terac t ions of prote ins wi th l ipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

C. Local izat ion of LPS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

D. Local iza t ion of phosphol ip ids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

E. Local iza t ion of ECA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87

F. Local iza t ion and topography of outer m e m b r a n e prote ins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87

1. In t roduc t ion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87

2. The major l ipoprote in . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88

3. O m p A prote in . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

4. Pept idoglycan-associa ted pore prote ins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90 5. Are matr ix (pore) prote ins associated wi th pep t idog lycan in vivo? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

G. The l ipid matr ix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93

1. Is the o u t e r m e m b r a n e a l ipid bi,layer? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93

2. Na ture of OM part icles and OM pits on the fracture faces of the outer m e m b r a n e observed with freeze-fracture

electron microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 3. Physical proper t ies of LPS and phosphol ip ids in the outer membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97

4. In terac t ions of prote ins with the l ipid matr ix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 5. Dis t r ibu t ion of outer membrane cons t i tuents over both monolayers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100

H. Molecular organiza t ion of the outer membrane of E n t e r o b a c t e r i a c e a e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 1. Outer m e m b r a n e of other Gram-nega t ive bacter ia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103

V. Future prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Acknowledgemen t s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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104

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I. Introduction

IA. Scope of this review

The cytosol of a bacterial cell is surrounded by a complex cell envelope which usually consists of a cytoplasmic membrane and a cell wall. Gram-posi-

tive and Gram-negative bacteria differ fundamen- tally with respect to the composition of their cell walls. One of the major differences is that Gram- negative cells contain an outer membrane, located at the outside of a monolayer of peptidoglycan. This outer membrane forms the physical and functional barrier between the inside of the cell and its

environment. After methods for the isolation of outer membranes became available, its composition, structure, function and biogenesis have been studied extensively in the last decade, especially in the enteric bacteria Escherichia coli and Salmonella typhimurium. Recently, a monograph [1] as well as several reviews have been published on the outer membrane in general [2,3] or on specific topics like genetics [4,5], functions [6-10] or biogenesis [5,7,11-16]. The present review will focus on the molecular architecture of the outer membrane and its constituents in relation to the membrane's functions.

IB. Ecological considerations relevant to structure and functioning of the outer membrane of Entero- bacteriaceae

The natural habitat of E. coli and other En- terobacteriaceae is the colon. From there these bacteria can reach the surface water where they can survive for quite a long time. Only specific classes of these bacteria can colonize or enter into other parts of the human or animal body, like cerebrospinal fluid, the blood stream and the urinary tract. Enterobacteriaceae constitute approx. 1% of all bacteria present in the gut, an anaerobic environment. Generation times are in the order of 10-20 h in contrast to only 20-60 rain under the usual laboratory conditions. In order to be able to compete with other micro-organisms (10u-10J2/g faeces) the bacterium should be able to take up nutrients effectively in order to remain in the gut. It also seems likely that the supply of nutrients varies both qualitatively and quantitatively, and it can be expected that the bacteria adapt continuously to such changes.

Bacteria can be attacked by other bacteria, by bacteriocins (proteins produced by bacteria which are toxic for other related bacteria) and by bacteriophages which bind to a specific receptor at the cell surface of a susceptible bacterium as the first step of a process that kills the cell. Survivors of such an attack are often mutants which lack the receptor for the bacteriocin or bacteriophage in question. As such receptors are often constituents of the outer membrane, bacteriocins and bacteriophages are excellent tools for studies on the outer membrane.

53

Donor cells of several bacterial species can transfer (part of) their genetic information to acceptor cells, the latter ones usually being related to the donor. Conjugative transfer, especially of plasmid DNA, usually is the means by which genetic information coding for production of toxins and for resistance against antibiotics, heavy metal ions or serum is passed from one cell to another, often even from one species to another. Because of the specificity of this conjugation process it is likely that again outer membrane receptors are involved.

Bacteria which live in the gut must somehow have a mechanism that protects the cytoplasmic membrane from the detergent-like action of bile salts, fatty acids and glycerides. Moreover, the gut content is rich in proteolytic and lipolytic enzymes and in glycosidases.

In order to be pathogenic, bacteria must have properties that enable them to adhere to eukaryotic cells a n d / o r invade tissue. Also, they must have mechanisms to resist the defence mechanism of the host (for a review, see Ref. 17). Among the most important pathogens are many Gram-negative bacteria e.g. Bordetella bronchiseptica, Bordetella pertussis, Campylobacter, E. coli, Erwinia amylovora and Erwinia carotovora, Haemophilis influenzae, Klebsiella, Neisseria gonorrhoeae, Neisseria meningitidis, Salmonella and Shigella. The relative contribution of Enterobacteriaceae as causative agents of infectious diseases in increasing, especially in elderly patients and in patients treated with broad spectrum antibiotics, corticosteroids and antimetabolites. It is often not realized by molecular biologists that E. coli is the leading cause of Gram-negative bacteremia in adults.

IC. General description of the cell envelope of Gram-negative bacteria

The cell envelope of Gram-negative bacteria has been the subject of a number of quite recent reviews [18-20]. This envelope (Fig. 1) consists of three essential layers, namely the cytoplasmic (or inner) membrane, the peptidoglycan (or murein) layer and the outer membrane. Although most soluble protein species are present in the cytosol, several of such protein species are located between the two membranes in a compartment known as

54

c ~ ~ta~elk~m

aody

Fig. 1. Features of the cell envelope of Gram-negative bacteria. The cytoplasm usually is surrounded by three layers: the cytoplasmic membrane, the peptidoglycan layer and the outer membrane. A few lipoprotein (LPP) molecules are drawn in the periplasmic space to indicate that these molecules are involved in anchoring the outer membrane to the peptidoglycan layer. The two membranes are connected by zones of adhesion (ZOA). Cells excrete part of their outer membrane into the medium as vesicles or blebs. A few ribosomes, as part of a membrane-bound polysome (MBP), are synthesizing proteins from which the exported (periplasmic and outer membrane) proteins are initially synthesized with a N-terminal signal sequence which is cleaved of during biogenesis. An A-layer or a capsular layer are sometimes present. The dimensions are not strictly drawn to scale. Especially the capsular layer can be much thicker than indicated. As the O-antigen (O-AG) and the flagellar body are very long, their ends have not been indicated. As the way of anchoring of pili is unknown, these structures have not been drawn. The basal body of the flagellum has been drawn sep- arately because the precise sites of connection between flagellar rings and envelope layers are not known. For further explana- lion see text.

the per ip lasmic space. The two membranes are in terconnected by so-called zones of adhesion. The outer membrane is not always the ou tmos t layer of the cell envelope as it is often covered with a ra ther amorphous capsular layer or with a so-called addi t iona l layer or A- layer consis t ing of a regular pa t t e rn of subunits , usually pro te ineous in nature. Final ly , appendices like flagella, f imbriae and pili are anchored in the cell envelope.

The cytoplasmic membrane conta ins phosphol ip - ids and prote ins in about equal amounts . It p lays a role in the t ranspor t of nutr ients , in oxidat ive phosphory la t ion , in the synthesis of phosphol ip ids , pept idoglycan , l ipopolysacchar ide and, via mem- b rane -bound polysomes, of per ip lasmic and mem- b rane proteins. Moreover , it is p robab ly involved in cell division and it serves as an anchor for D N A , at least dur ing repl icat ion.

The peptidoglycan layer consists of a network in which linear amino sugar chains, conta in ing alter-

nat ing residues of N-ace ty lg lucosamine and N- ace ty lmuramic acid, are covalent ly l inked to each other via crossl inks that can be formed between te t rapept ides which are a t tached to the N- ace ty lmuramic acid residues (see Ref. 21 for a review on pep t idog lycan structure). In Gram-negative bacter ia a monolayer of pep t idoglycan sur- rounds the cy toplasmic membrane . The r igidity of the pep t idog lycan layer enables the cell to with- s tand the osmot ic pressure of approx. 3.5 a tm [22] of the cytoplasm. Degrada t ion of pep t idog lycan

e.g. by lysozyme, results in lysis of the cell as a consequence of heavy swelling caused by the uptake of water through the cytoplasmic membrane into the cytosol. Disrupture of the cytoplasmic membrane can be prevented if pep t idog lycan is degraded under hyper tonic condi t ions e.g. in the presence of 10% sucrose. As the rods then loose their shape and round up to osmot ical ly fragile spheroplas ts , pep t idog lycan is p robab ly (co-)responsible for the rod shape as well. Consis tent with this assumpt ion is the observat ion that isola ted pep t idog lycan has the same shape as the cells f rom which it has been isolated.

The outer membrane contains, in addi t ion to phospho l ip id and protein, LPS as a major const i tuent. The membrane is covalent ly a t tached to the pep t idog lycan layer via a l ipoprotein. The m e m b r a n e prevents leakage of per iplasmic proteins. Moreover, in Enterobacteriaceae it has to protect the content of the cell against the a t tack by harmful agents like bile salts and enzymes. The extent of this protect ive function is very impressive as E. coli can grow in the presence of as much as 5% of the s trong ionic detergent SDS. As phospho l ip id bi layers are very sensitive to this detergent (erythrocytes lyse in the presence of as little as 0.001% SDS (Bergmans, H.E.N., Overbeeke, N. and Van Scharrenburg, G., personal communication)), this SDS resistance indicates that the outer membrane either is not a phospho l ip id bi layer or that its bi layer s tructure is very well shielded from the medium.

The periplasmic space is located between the two membranes . It has been repor ted to comprise as much as 20--40% of the total cell volume [22] a l though much lower values have also been repor ted [9]. The space contains prote ins [23] and ol igosacchar ides [24]. The per ip lasmic proteins

55

comprise approx. 4% of the total cell protein [23]. With respect to their functions three classes of periplasmic proteins can be distinguished [23,25]. Proteins with a catabolic function (e.g. 5'- nucleotidase and alkaline phosphatase) convert solutes for which no transport system exists to a form that can be transported through the cytoplasmic membrane. Another class of periplasmic proteins are the binding proteins which have affinity for nutrients like sugars, amino acids or ions. It has been established in a number of cases that these proteins are essential for transport of the solute in question. A third class of periplasmic proteins is involved in the degradation or modification of harmful components such as antibiotics and heavy metals.

Periplasmic membrane derived oligosaccharides [24,26,27], comprising about 1% of the dry weight of the cell, are a closely related family of highly branched molecules, containing about nine residues of glucose as the sole sugar. They are vari- ously substituted with sn-glycerolphosphate and phosphatidylethanolamine residues derived from the membrane phospholipids. Some species of the oligosaccharides also contain P-succinyl esters, ad- ding to their net negative charge. Membrane-derived oligosaccharides may play an important role in the osmoregulation of the Gram-negative cells as cells grown in medium with low osmolarity synthesize 16 times more of these substances than they do under conditions of high osmolarity [28].

Elegant experiments of Stock et al. [22] have shown that there is a Donnan equilibrium over the outer membrane, and that the periplasm and cytoplasm are isosmotic. For cells in minimal medium, the osmotic strength of the cell interior was estimated to be approx. 300 mosM [22].

'Zones of adhesion" [ 12] are electron microscopi- cally visible contact sites between cytoplasmic and outer membrane which become apparent only when the membranes are separated by plasmolysis [29]. About 200-400 of these sites are present per cell, they measure 20-30 nm across and cover about 5% of the membrane surface [12]. Several lines of evidence indicate a physiological role for these 'zones of adhesion'. Firstly, in phage-in- fected cells empty phage heads are found opposite adhesion zones, strongly suggesting that they are involved in the penetration of phage DNA into the

cell [29,30]. Secondly, these zones have been implicated in the translocation of newly synthesized LPS [12,35,36] in the synthesis [32,33] and translocation [34] of (some) outer membrane proteins and in the production of sex pili [12].

Assuming that these 'zones of adhesion' are stable structures several investigators have undertaken attempts to purify these domains, but consistent data on their composition have not been obtained so far. Recently a fraction, presumed to consist of these domains, was reported to be enriched in phospholipase activities [31].

An alternative for permanent 'zones of adhesion' is that these zones exist only temporarily. Basically this idea comes from Witholt and co- workers [32,33], who suggested that these 'zones of adhesion' might be sites of outer membrane protein synthesis generated by the fact that the protein synthesizing machinery is producing an outer membrane protein. The consequence would be that these zones exist no longer when the protein chain has been completed.

Considering biogenesis of outer membrane proteins at the membrane level at specialized temporary domains in the cytoplasmic a n d / o r outer membrane one can basically propose two mechanisms, both of which are based on the presence of non-bilayer lipids. In the first model a protein is exported via a vesicle which blebs off from the cytoplasmic membrane and fuses with the outer membrane. According to the second mechanism the protein diffuses through a temporary connection between the two membranes. The observation that E. coli membrane phospholipids [37] and also LPS [162,176] have the tendency to form non-bilayer structures is consistent with these models since these non-bilayer phases, especially the hexagonal II phase, are supposed to represent intermediate stages in fusion processes [38]. This hypothesis is strongly supported by freeze-fracture experiments in model membrane systems [39,40], including those con ta in ing phospha t idy l - ethanolamine [41], the major phospholipid of E. coli. For reviews on this subject the reader is referred to Refs. 42, 43 and 44.

Based on these considerations we propose three models for functional meta-stable connections between cytoplasmic and outer membrane (Fig. 2). It should be noted that direct evidence in support of

56

A Lll:~T . . . . b c d

e f l

B

OM

CM

C

a ,o,

, Id) l~ OM ~ { ~t~

(allL I(a)

Fig. 2. Models for connections between cytoplasmic (CM) and outer membrane (OM). A: Vesicle-mediated fusion model. LPS (L) and outer membrane proteins (P) are synthesized at the cytoplasmic membrane (a). Blebbing starts in certain special domains of the cytoplasmic membrane, possibly due to the presence of a high concentration of LPS molecules or due to the presence or formation of protein-LPS complexes. The bleb pinches off (b) and subsequently fuses with the same membrane (c), e.g. by a mechanism illustrated in Fig. 2B or C. in which the lipids phosphatidylethanolamine, cardiolipin and/or LPS, which are known to be able to participate in non-bilayer phases, may be involved. This transient state is followed by the release of a vesicle into the periplasm (d). The vesicle fuses with the outer membrane (e), resulting in the insertion of LPS and protein, or of a complex of these molecules, in the outer membrane in the correct orientation (f, g). B and C: Direct fusion models. A LPS molecule in the inner monolayer of the cytoplasmic membrane (a) is synthesized in or diffuses to (b) a special domain in the cytoplasmic membrane (see above). Due

the se m o d e l s is l ack ing , b u t t h a t they are r a t h e r

m e a n t to s t i m u l a t e i n v e s t i g a t o r s to tes t the a t t r ac -

t ive pos s ib i l i t y of t e m p o r a r y a d h e s i o n sites. T h e

o b s e r v e d role of the "zones of a d h e s i o n ' in the

t r a n s l o c a t i o n of LPS a n d of at leas t s o m e o u t e r

m e m b r a n e p r o t e i n s has b e e n i n c o r p o r a t e d in these

mode l s . M o r e o v e r , t he pos s ib i l i t y t h a t o u t e r m e m -

b r a n e p r o t e i n s a n d LPS, w h i c h o f t e n fo rm func-

t i o n a l c o m p l e x e s (see s ec t i on IV), a re t r a n s l o c a t e d

as a c o m p l e x or a re even i n v o l v e d in the i n d u c t i o n

of fus ion , c a n eas i ly b e i n c o r p o r a t e d in the m o d -

els.

T h e f i rs t m o d e l (Fig. 2A) is b a s e d o n m o d e l s

p u b l i s h e d r e c e n t l y to e x p l a i n t r a n s l o c a t i o n of LPS

[45] a n d of o u t e r m e m b r a n e p r o t e i n s [15,16[, in

w h i c h it is p r o p o s e d t h a t a vesicle is r e l ea sed f r o m

the c y t o p l a s m i c m e m b r a n e b y fus ion a n d subse -

q u e n t l y fuses w i th the o u t e r m e m b r a n e . T h i s m o d e l

c a n be e x t e n d e d in t h a t the s i te of t r a n s l o c a t i o n is

s u p p o s e d to c o r r e s p o n d w i th a ' z o n e of a d h e s i o n '

(F ig . 2A). B l e b b i n g s t a r t s at a s i te in the cy to-

p l a s m i c m e m b r a n e , w h i c h has a spec ia l c o m p o s i -

t i on (e.g. r i ch in LPS or c e r t a i n p h o s p h o l i p i d s ) or

s t r u c t u r e (e.g. a s t r o n g l y a s y m m e t r i c r eg ion d u e to

t he p r e s e n c e of a la rge n u m b e r of LPS m o l e c u l e s

in the i n n e r m o n o l a y e r or to the p r e s e n c e of or

f o r m a t i o n of c o m p l e x e s of o u t e r m e m b r a n e p ro -

t e in s a n d LPS). A vesic le is f o r m e d a n d subse -

q u e n t l y fuses w i th the o u t e r m e m b r a n e , p e r h a p s at

spec ia l sites. M o l e c u l a r m o d e l s for fus ion a n d

to the special structure in the site of fusion, in which a lipidic particle may be absent (Fig. 2B) or present (Fig. 2C), the LPS molecule reaches, by a flip-flop mechanism, first the area where the outer monolayer of the cytoplasmic membrane is connected with the inner monolayer of the outer membrane (c) and subsequently the outer monolayer of the outer membrane (d). It is conceivable that outer membrane proteins either are synthesized at arbitrary sites in the cytoplasmic membrane or at such adhesion zones and that they form complexes with LPS molecules before, during or after fusion. As no clear choice can be made between all these possibilities, outer membrane proteins have only been drawn at their final location in the outer membrane. Finally, it should be noted that all three models allow, but not require, the translocation of phospholipid to the outer monolayer of the outer membrane (see Figs. 2B and 2C). It has clearly been established that the outer leaflet of E. coh contains no or hardly any phospholipid (see section IV), but it has been argued that fusion of bacterial cells with phospholipid vesicles is easier to envisage if some phospholipid is present in the outer monolayer (see subsection IVG-1/.

57

fission of such vesicles are shown in Figs. 2B and C. The translocation of LPS with its hydrophilic sugar chain is well-explained by this vesicle-mediated translocation model. However, such a long sugar chain will require vesicles with a rather large diameter, which can be estimated to be at least 40 nm, unless the O-antigen sugar chains are somehow organized close to the bilayer during this process. This large diameter would require, at least locally, a thick periplasmic space. It can be argued that it also would require rather large gaps in the peptidoglycan layer, which have never been detected by electron microscopy. However, if growth of the vesicle occurs outside the peptidoglycan layer, much smaller - perhaps undetectible - gaps would he sufficient. It should be noted that the occurrence of gaps in the peptidoglycan is not unlikely as the total amount of peptidoglycan in the cell is only sufficient to cover half of the cell surface. The model shown in Fig. 2A also implies a small leakage of cytoplasmic constituents into the medium. Vesicle-mediated excretion of soluble constituents of the cell has been described re- peatedly, e.g. during normal growth E. coli releases outer membrane vesicles or blebs, corresponding with approx. 5% of the total outer membrane, into the medium [46-49]. An offensive function has been proposed for these vesicles in the delivery of heat-labile enterotoxin by E. coli cells to intestinal epithelial cells [50].

Two other models for hypothetical recta-stable connections between the two membranes are shown in Figs. 2B and 2C, both of which propose a direct fusion between the membranes. Whereas in Fig. 2B no special structure is proposed within the fusion site, an 'inverted' micelle is proposed in Fig. 2C. Although especially the translocation of LPS molecules with their hydrophilic, sometimes very long, sugar chains through flip-flop, even in the special environment of the fusion site is some- what hard to imagine, it should be noted that it has been established that phospholipid molecules do use such structures to cross membranes [41]. Moreover, evidence that the long O-antigen chains of LPS can flip-flop over a membrane comes from the observation that these molecules redistribute very rapidly over both membrane monolayers upon

incubation of isolated outer membranes with lysozyme at 37°C but not at 0°C [543].

It is very likely that both the blebs shown in Fig. 2A and the connections between the two membranes drawn in Figs. 2B and C are extremely unstable and that, unless they are somehow stabi- lized (e.g. by protein synthesis on membrane-bound ribosomes) they cannot be detected by electron microscopy. Consequently, the detection of 'zones of adhesion' in thin sections could be due to artificial blebbing or fusion e.g. at sites in the cytoplasmic (or outer) membrane where a blebbing or fusion process is going to start.

Flagella (H-antigen), fimbriae and pili are ap- pendages, consisting of protein subunits. Flagella are responsible for the cells motility. They are connected with all three layers of the cell envelope [51]. Fimbriae are smaller and more rigid than flagella [52,53]. They occur in many varieties and several types have been implicated in adhesion to (often specific) eukaryotic cells [54,55]. A special type of fimbriae, called pili, is essential for the adhesion of bacterial donor cells to acceptor cells, the first step in bacterial conjugation [6,8].

A capsular layer (K-antigen) usually consisting of negatively charged polysaccharides [56,57], can be present outside the outer membrane. The recent discovery of lipid constituents in the K92 capsule lead the authors to suggest that the lipid portion enables the capsule to anchor in the outer membrane [58]. The presence of a capsule sometimes enhances the resistance of the cell against phago- cytosis and against the bactericidal action of com- plement.

Additional layers [59,60] or A-layers are surface structures consisting of regularly arranged subunits of usually one protein species. They are located outside the outer membrane and have been discovered in several bacterial species like Acin- etobacter [61,62], Azotobacter vinelandii [63], Campylobacter fetus [64], Spirillurn [65-68], Treponema and, more recently, in Aeromonas salrnonicida [69-71]. It has been proposed that such layers play a role in protecting the cell against a harmful environment [60,72] or that they are involved in adherence to specific tissues [73].

58

II. Methods for the isolation of outer membranes

IIA. E. coli and S. typhimurium

IIA-1. Isolation of peptidoglycan-less outer membranes after spheroplast formation.

If necessary, cells can first be treated in an omnimixer in order to remove flagella, pili and capsular material, a procedure which does not result in cell breakage [74]. One of the most critical steps in most procedures used for separating cytoplasmic and outer membranes consists of converting cells to spheroplasts. These are then lysed to yield outer and cytoplasmic membrane vesicles [75-77]. Outer membranes can be isolated on density gradients [75-79], by electrophoretic techniques [80-82] or by aggregation at pH 5, followed by centrifugation [83]. Lysozyme, which cosep- arates with the membranes, can be removed by washing with 0.2 M KC1 [76]. Outer membranes, isolated from spheroplasts, have a buoyant density of 1.22 _+ 0.01. They contain approx. 60% of the protein, 50% of the phospholipid and 90% of the LPS of the cell envelope [76].

Birdsell and Cota-Robles [84] have described the basic procedure for the production of spheroplasts which can serve as the basis of a good membrane separation. Slight modifications of this procedure have extensively been described by Osborn et al. [76] and Witholt et al. [77]. Basically, the cells are plasmolyzed with 0.5 M sucrose, lysozyme is added, and the outer membrane is made permeable for this peptidoglycan-degrading enzyme by treatment with EDTA. The procedure converts rod-shaped cells into osmotically sensitive spheroplasts i.e. the cytosol surrounded by an intact cytoplasmic membrane with outer membrane structures attached to it at one or only a few sites [84].

Miura and Mizushima [75,78] were the first investigators who successfully separated cytoplasmic and outer membranes. In our hands the best results are obtained with the procedure of Osborn et al. [76] who slightly modified the procedure of Birdsell and Cota Robles [84] for the preparation of spheroplasts. Moreover, the Osborn procedure can completely be carried out at 4°C and does not result in significant release of LPS

from the outer membrane which usually accompa- nies EDTA treatment [85]. More recently Witholt et al. [77] introduced a similar procedure which. however, has the advantage that it is applicable to laboratory strains of E. coli grown under various conditions [77]. Also, it turned out to be the procedure of choice for the isolation of outer membranes from clinical strains of E. coil (M. Acht- man, personal communication).

llA-2. Isolation of outer membrane-peptidoglycan complexes

Due to the pioneering work of Schnaitman [86], the outer membrane can be isolated complexed to the peptidoglycan layer. The method involves cell disruption in a French pressure cell followed by membrane separation on a sucrose density gradient. Slightly modified versions of this procedure have been published by Koplow and Goldfine [87] and Smit et al. [88]. Neither the original procedure nor any of the mentioned modifications could be applied successfully in our laboratory but the introduction of another modification [37] resulted in a good separation of the two membranes. For structural studies the use of outer membrane- peptidoglycan complexes has several advantages over the use of outer membranes prepared from EDTA-lysozyme spheroplasts. (i) The use of EDTA, a prerequisite in the latter method as it is required to make the outer membrane permeable to lysozyme, is avoided. (ii) Removal of the peptidoglycan-layer by lysozyme results in reorganization of the LPS component [89]. (iii) Cy- toplasmic and outer membranes of heptose-less mutants cannot be separated using the EDTA- lysozyme method [76] but the French press method gives good results [87,88]. The most likely explanation for the observed failure in the former case is that the buoyant density of the peptidoglycan-less outer membrane of heptose-less mutants is similar to that of the cytoplasmic membrane, presumably due to the loss of sugars as a direct result of the mutation (see Fig. 3), as well as due to the loss of protein and an increase in phospholipid content as an indirect result of the mutation [87,90-96]. The fact that the presence of peptidoglycan increases the density of the outer membrane [37,88] is the basis for the success of the separation method developed by Schnaitman. A possible disad-

vantage of the latter method is that phospholipids are degraded by the action of an outer membrane-bound phospholipase AI (see subsection IIIE-2). This disadvantage can be overcome by the use of a mutant lacking this activity [37].

11,4-3. 'Differential membrane solubilization using detergents

It is a rather common viewpoint that treatment of isolated cell envelopes with certain detergents can selectively solubilize the cytoplasmic membrane while leaving the outer membrane intact. It is our view that, at best, the proposed selectivity has been proven for the protein component of the membranes. The effect of Triton X-100 on the solubilization of membrane fractions has been studied in detail [97]. Extraction of cell envelopes with 2% Triton X-100 in the presence of mag- nesium ions results in complete solubilization of the cytoplasmic membrane. Morphologically the outer membrane seems to remain intact. However, chemical analyses showed that this 'Triton-insoluble cell wall' contains only about half of the LPS and one third of the phospholipid of the outer membrane but that all outer membrane proteins are still present [97]. Filip et al. [98] have studied the effect of a large number of detergents on the integrity of the E. coli membranes. From their data it can be concluded that, in the absence of mag- nesium ions, treatment with 0.5% sodium lauryl sarcosinate (Sarkosyl) results in solubilization of the cytoplasmic membrane proteins and phospholipids. With respect to the outer membrane the authors do not provide solid quantitative data on the fate of the various constituents. They have not analysed the fate of LPS whereas their data suggest that treatment with Sarkosyl results in an increase of the buoyant density of the outer membrane, suggesting loss of phospholipid.

11.4-4. Membrane separation based on charge differences of vesicles

Using the property that outer membrane vesicles have a larger negative charge than cytoplasmic membrane vesicles, they can be separated in a preparative particle electropherograph [81,99[ or by sucrose gradient electrophoresis [82].

59

liB. Other organisms

Procedures developed for the separation of the membranes of E. coli and S. typhimurium have been applied successfully on other Gram-negative bacteria, usually after some modification of the procedure. Examples are Acinetobacter sp. [100], Campylobacter fetus [ l01 ], Caulobacter crescentus [102,103], Chlamydia trachomatis [104], Chro- matium vinosum [105], Haemophilus influenzae [106,107], Klebsiella aerogenes [108], Moraxella nonliquefaciens [109], Myxococcus xanthus [ll0], Neisseria meningitidis [111], Neisseria gonorrhoeae [112,113], Proteus mirabilis [114], Pseudomonas aeruginosa [ 115-117], Rickettsia prowazeki [ 118], Selenomonas ruminantium [119], Serratia marcescens [120], Yersinia pest& Y. enterocolitica and Y. pseudotuberculosis [121]. It should be noted that methods based on differential solubilization should be thoroughly checked by the use of methods based on a physical separation as it has been found that extraction of H. influenzae membrane with Triton X-100-Mg 2÷ results in the extraction of some major outer membrane protein species (Van Alphen, L., unpublished data). A method often used for the isolation of certain outer membrane fractions consists of the extraction of cells with approx. 0.2 M LiC1 (as was used for N. meningitidis, Ref. 111), Li-acetate (as was used for N. gonorrhoeae, Refs. 122 and 123) or 0.5 M NaC1 followed by 0.5 M sucrose (used for Gram-negative marine bacteria, Refs. 124 and 125). This method is not successful if applied to Enterob- acteriaceae or to H. influenzae (Van Alphen, L., unpublished data).

IlL Individual constituents of the outer membrane

IliA. Composition of the outer membranes

As determined by electron microscopy the outer membrane, like the cytoplasmic membrane, has a thickness of 7.5 nm [59]. It contains protein (9-12% of the cellular protein), LPS and phospholipid as its major constituents. Enterobacterial Common Antigen (ECA) is a minor component (0.2% of the cellular dry weight).

A comparison of the data from various laboratories on the weight ratio's of the three major

60

constituents [76,88,92,93,114,126-129] shows that these differ enormously, even if one takes into account the influence of the length of the sugar chain of the LPS (see subsection IIIC). For example, in the case of E. coli weight ratio's for outer membrane phospholipid : LPS : protein of approx. 1 : 1 : 1 [129] and 1 : 1:6 [126] have been reported. Another striking example is that, despite the fact that the groups of both Nikaido and Gmeiner have undertaken extremely serious attempts to quantify the various outer membrane constituents, the former group reports that the number of LPS molecules per unit surface area is not influenced by a deep rough mutation [88], whereas the latter group claims a 4-fold increase of the relative amount of LPS in the outer membrane of the mutant compared with that of the parental strain [92,93]. As in both laboratories LPS was determined by assaying the content of 3-hydroxytetradecanoic acid, differences in the methods used are not a likely explanation in this case. In contrast, the method used for protein assay can influence the data as it has been shown by Schweizer et ai. [129] that too high values are obtained for the protein content of outer membrane preparations with the method of Lowry et al. [130]. Also, data from our laboratory show that the method of Lowry et al. yields values that are 2-3-fold higher than those obtained by scanning of stained polyacrylamide gels [127]. In addition, it is conceivable that some of the published differences in the composition of the outer membrane can partly be caused by factors like the bacterial species, the particular strain chosen, the growth conditions and the growth phase. In conclusion, in order to be able to perform detailed calculations on outer membrane composition a thorough evaluation of assay procedures is required to establish which data can be used. Moreover, details on experimental growth conditions should be standardized carefully in order to allow the comparison of data from different laboratories.

IIIB. Phospholipid

Essentially all phospholipids of E. coli are located in the cell envelope [131]. Phosphatidy- lethanolamine is the major species whereas substantial amounts of phosphatidylglycerol and di-

phosphatidylglycerol (or cardiolipin) are found. Excellent reviews on various aspects of phospholipids of Gram-negative bacteria have been published [ 131-135].

Relatively little attention has been paid to the distribution of the phospholipids over the two individual membranes. Osborn et al. [76,136] reported that the two membranes of S. typhimurium contain equal amounts of phospholipid and that the outer membrane is enriched in phosphatidylethanolamine whereas the cytoplasmic membrane is enriched in the other two major species. The lipid composition of the two membranes of E. coli has extensively been studied in our laboratory [137]. Consistent with the results of Osborn et al. [76] we observed that the outer membrane is enriched in phosphatidylethanolamine, which -~ in contrast to the data reported for S. typhimurium [76] - often represented over 90% of the outer membrane phospholipid ]137]. Similar high amounts of phosphatidylethanolamine in E. coli outer membranes have been found by R.M. Bell (cited in Ref. 138). To our knowledge the only data conflicting with the above mentioned distribution are those of White et al. [81] who found considerably more phosphatidylethanolamine in the cytoplasmic than in the outer membrane. This is probably caused by a preferential degradation of the outer membrane phosphatidylethanolamine by phospholipase and lysophospholipase activities located in this membrane (see subsection IIIE-2). Consistent with this explanation is the high percentage of lysophosphatidylethanolamine found by these authors in the outer membrane, Only trace amounts of the latter phospholipid were found in other laboratories [76,137]. The relative abundance of phosphatidylethanolamine in the outer membrane was found in various strains under conditions in which the fatty acid composition was altered by the growth temperature, by mutation or by supplementation of the growth medium with fatty acids [137]. The reason for the enrichment of the outer membrane with phosphatidylethanolamine might be that it forms stable bilayers with LPS [139]. An alternative explanation, namely the association of major outer membrane proteins with this phospholipid, can be excluded as a similar enrichment is found in mutants lacking the three major outer membrane proteins OmpA protein,

61

OmpC protein and OmpF protein (Van Alphen, L. and Lugtenberg, B., unpublished data). The enrichment is remarkable since a fast exchange between the phospholipids of the two membranes was reported by Jones and Osborn who fused lipid vesicles with intact cells of S. typhimurium [140]. Vesicle lipids consisting of total Salmonella phospholipid are recovered in both cytoplasmic and outer membranes without clear preference for either of the membranes. Moreover, these authors showed that, after fusion, phosphatidylserine from vesicles is converted to phosphatidylethanolamine which can be recovered from both membranes. Since the enzyme responsible for this conversion is located in the cytoplasmic membrane, exchange of phospholipids must occur in the membranes in both directions [136,140]. Application of the fusion technique also enabled the authors to see whether lipids, which are not normal constituents of Enterobacteriaceae can be incorporated into their membranes. They could show that phosphati- dylcholine was incorporated into both membranes without a significant preference. Similar experiments using cholesterol oleate suggested that this lipid showed preferential accumulation in the cytoplasmic membrane [ 136].

~ c ~ 53--{ co,o - - } - t o ........ 1

I - R e LPS I - - RO2LPS - - I

RO 1 LPS - I

- - Rc LPS I

Rb LPS q

Ra LPS - - d

F S LPS I

Fig. 3. Schematic representation of the chemical structure of LPS from S. typhimuriurn. Wavy lines represent long chain fatty acids (C8-C16). The number of repeating O-antigen units n is variable (see text). Wild-type cells synthesize the entire structure (S-LPS) whereas various mutant strains produce the chemotypes Ra through Re LPS. The structures synthesized by so-called deep-rough mutants are rather glycolipids than lipopolysaccharides. Note the large number of charged residues in the lipid A-core region of the molecule. The phosphate residues on the glucosamines are sometimes substituted (see Fig. 4). EA, ethanolamine.

The major fatty acids in E. coli phospholipids are palmitic acid (C16 : 0), palmitoleic acid (C16 : 1) and cis-vaccenic acid (C18:1) or their cyclopropane derivatives [131]. A slight but significant enrichment of the outer membrane with saturated fatty acids is found whereas the cytoplasmic membrane is enriched in unsaturated and cyclopropane fatty acids [81,137]. This enrichment is independent of changes in the fatty acid composition by mutation or by altered growth conditions [137]. Detailed studies have shown that phosphatidylethanolamine of the outer membrane contains more saturated fatty acids [137] and less molecular species with two unsaturated fatty acids [141] than phosphatidylethanolamine of the cytoplasmic membrane.

IIIC. Lipopolysaccharide (LPS)

IHC-1. Introduction LPS, which is characteristic for Gram-negative

bacteria, is an amphipathic molecule with a hydrophobic part, called lipid A, and a hydrophilic, often branched, sugar chain. The hydrophilic part consists of an oligosaccharide core which usually is substituted by the O-antigen, the latter being a polymer consisting of repeating carbohydrate units. A schematic representation of the general structure of LPS is given in Fig. 3. Colonies of strains with or without the O-antigen often have a S(mooth) or R(ough) appearance respectively. Therefore one often speaks of S and R strains and also of S and R types of LPS. Among Entero- bacteriaceae the structure of the lipid A core region is rather well conserved but that of the O-antigen has been subject to extreme evolutionary changes. Many reviews have been written on LPS in genera l [142,143], on its chemica l [56,142,145-150], physical [144] and biological [56,142,145,150,151 ] properties and on its genetics [ i 52,153] and synthesis [45,154,155]. Therefore we will only briefly outline some general aspects of LPS.

IIIC-2. Methods of isolation and purification The standard method for the isolation of pure

LPS has long been the phenol-water method, i.e. ,treatment of dried bacteria with 45% phenol for 5 min at 68°C [156,157]. The water phase obtained

62

after cooling contains nucleic acids as the main contaminants, which can be removed by ultra- centrifugation. Even better results were obtained when cells as the starting material were replaced by cell envelopes [157a]. The metal ions Na ~ , Mg 2+ and Ca 2+ as well as a number of amines such as ethanolamine, putrescine, cadaverine, spermidine and spermine are often found in LPS preparations in variable amounts, depending on growth and isolation conditions [158]. They have a profound influence on the physico-chemical properties of LPS and can partly be removed by electrodialysis [158].

The phenol-water method is applicable for S- LPS and Ra and Rb LPS. Since LPS is isolated in low yields when the phenol-water method is applied to Rc, Rd and Re mutant bacteria (see Fig. 3) a more hydrophobic solvent was developed which is adequate for the extraction of R-LPS [159]. This very mild procedure involves extraction of dried bacteria at room temperature with a mix- ture of 90% phenol, chloroform and petroleum ether (PCP method). Other methods include extraction of bacteria with either EDTA [160,161] or aqueous butanol [160,162] procedures which have not been extensively tested for other bacteria than E. coli.

Isolated LPS forms multimeric states. It can have the morphological appearance of droplets, long rods, unilamellar bilayer vesicles, stacked lamellae, long ribbons, flat sheets or a doughnut [ 144,162,163]. Under certain conditions particles and pits can be seen on fracture planes [162]. The morphological appearance depends on the length of the sugar chain [144], the degree of purity, the composition of the solution used for hydration, the presence of Ca 2 + and on whether the structure is studied at temperatures above or below the phase transition temperature [162]. Van der Waals interactions between the fatty acyl chains of lipid A moieties are the basis for the large structures. The variety of macromolecular organizations are probably caused by ionic interactions due to salt bridges between a variety of negatively (phosphate, KDO and uronic sugars) and positively (glucosamine, 4-amino-arabinose, ethanolamine and amino sugars) charged residues. Therefore, the 'solubility' of LPS can be increased by both disturbing the salt bridges, by electrodialysis or by the addition

of EDTA [117], as well as by the weakening of the hydrophobic interactions, by the addition of detergents [162,164] or triethylamine [142,158].

11IC-3. Chemical structure of lipopolysaccharides The LPS of S. (vphimurium has been studied

most extensively. Next to chemical methods the availability of mutants has largely contributed to the elucidation of LPS structures. Lipopolysac- charides of wild-type and mutant strains have been classified into chemotypes based on the content of their sugar constituents (see Fig. 3).

Lipid A can be obtained by mild acid hydrolysis of LPS. Lipid A of Enterobacteriaceae is a glycolipid which contains a/3-(1 ~ 6) linked glucosamine disaccharide unit which carries a phosphate residue in position 1 and a phosphate or pyrophosphate residue in position 4' and also approx. six fatty acyl chains (Refs. 169 and 165- 168, see also Fig. 4), two of which are amide-bound. Various ligands (glucosamine, ethanolamine, ions) can be attached non-covalently. Among the fatty acids /3-hydroxy fatty acids are abundant. Un- saturated fatty acids are hardly present and most of the fatty acids are relatively short (C8-C14) compared with the fatty acids found in the phospholipids. The amide-linked/3-hydroxy fatty acids are specific for LPS. In Enterobacteriaceae fi-hydroxy tetradecanoic acid is the major amide-linked fatty acid. Like dodecanoic and tetradecanoic acid it can also be ester-linked. Tetradecanoic acid also occurs as part of a branched fatty acid when it is esterified to the hydroxyl moiety of the/3-hydroxy fatty acids [142,165,168,170,171]. It should be noted that also other types of lipid A backbones exist for which those of Chromobacterium and Rhodopseudomonas eiridis and standard examples [142,146].

Lipid A preparations are heterogeneous [166,172,173]. It appears that non-equimolar amounts of 4-amino arabinose [174] and of the esterbound phosphate residues at C4' [166,167, 175] as well as a disproportionate distribution of ester-bound fatty acids [168] contribute to this heterogeneity. Moreover, growth conditions have been shown to influence the fatty acid pattern of lipid A [170,596].

It has long been thought that the phosphate residues of lipid A were involved in the formation

< i

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0 R R R

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Fig. 4. Proposed structure of lipid A from S. minnesota. From Refs. 165 and 165a. This type of lipid A is probably general for Enterobacteriaceae. Number of fatty acid carbon atoms: a = 14, b = 12, C = 16. The exact position of amide-bound and esterbound 3-acyloxytetradecanoic and 3-hydroxytetradecanoic acids has not yet been detertnined. It is possible that also nonacylated 3-OH-14 : 0 as well as trace amounts of 3-O(10 : 0)- 14:0 and 3-O(12:0)-12:0 are amide-bound. The sum of esterbound 3-O(14: 0)- 14 : 0 and 3-O(2-OH- 14 : 0)- 14 : 0 equals approx. 1 mol /mol lipid A. Since three hydroxyl groups are available at the lipid A backbone and since up to now only two Ooacyl groups have been detected it is possible that one glucosamine hydroxyl group is not substituted (R ~ = H). The hydroxyl group in position C-3' represents the attachment site of the polysaccharide portion. Residues 4-amino-4-deoxy-L- arabinose and phosphorylethanolamine are not present in molar amounts as indicated by dotted bonds.

of cross-links between monomers, thus forming trimers [176]. However such cross-links were not detected with 31p-NMR by MOhlradt et al. [165]. We therefore assume that the LPS molecule is monomeric.

The LPS core of Enterobacteriaceae contains the unique sugars 3-deoxy-D-manno-octulosonic acid (KDO) and L-glycero-D-mannoheptose, both of which are practically LPS-specific. In addition, it contains a number of more common sugars like glucose, galactose and N-acetyl-D-glucosamine [ 142,147,150]. Numerous mutants with defects in the structure of LPS have been isolated, usually by selection for resistance towards a certain bacteriophage or towards the antibiotic polymyxin [177,178]. Mutants completely missing KDO can be isolated as conditionally lethal [172,179,180]. Ra mutants are defective in the biosynthesis of

63

O-antigen and have a complete core whereas Re mutants lack all Hep residues (Fig. 3). Prehm et al. [181] have analyzed the core structure of E. coli K-12 LPS in detail (Fig. 5). The core of this strain, which lacks the O-antigen, is heterogenous in that several core structures are found with various de- grees of completion. The structure of the core region of strains of Enterobaceriareae is similar. However as in quite a few other Gram-negative bacteria KDO or L-glycero-D-mannoheptose or both are lacking, they must have different core structures. Examples are Acinetobacter sp., A na£vs- tis nidulans, Bacteroides spp. [142], Haemophilus influenzae [182] Moraxella sp., Pseudomonas spp., Spirillum serpens and Vibrio cholerae [142].

The O-antigen can consist of more than 40 repeating units containing 3 to 6 sugar residues. The number of repeating units can vary, even in a culture of one strain [183-185], from none to more than 40, thereby providing the cell with the op- portunity for subtle variations in the molecular make up at different sites at its surface. The structure of the subunit of the O-antigen shows extreme diversity, even within a single genus like E. coli or S. typhimurium. This property is used in O-sero- typing, an immunological method used to identify sub-strains of one species in great detail [56]. As thus could be expected, a large diversity of sugars has been found in the repeating units of the O-antigen. These include neutral sugars, amino sugars and uronic acids. Moreover sugars can be substituted with O-acetyl groups, phosphate, amino acids or even ethanolamine triphosphate. The

C)4 0 Cl4

IC Z O ~ ' ,~

G , c N A c - ~ , c ~ G , ~ O , c ~ H ~ 0 _ , . . Kt~Cl o. f16 -,:,- f ( e ~'~- ,.

Gal Hep Rha EA

~te'p7

Fig. 5. Tentative structure of E. coli K-12 LPS composed from data of Refs. 142, 165, 171 and 181. Ions and other non-covalently attached constituents are not included. Amide and ester bonds are indicated by -N- and -O- respectively. The exact location of the fatty acid residues is not known.

64

O-antigen therefore can contribute considerably to the net surface charge of the bacterial cell. The O-antigen is not present in all strains. For instance such a structure is lacking in the E. coli laboratory strains K-12 [181], B [186] and C [187] and also in some strains with for instance serotype O14 [56]. Also, within a certain strain the presence of the O-antigen can depend on the growth condition [188] whereas the presence of certain prophages in the chromosome can drastically alter the structure and composition of the O-antigen, a phenomenon known as antigenic conversion [189,190]. Also the presence of plasmids [191,192] can alter the composition of the LPS. The structures of only a few O-antigens have been established [56].

IIIC-4. Effects of polymyxin, EDTA and divalent cations

Polymyxin B is a polycationic amphipathic antibiotic which is bacteriacidal to most Gram-negative bacteria and binds to their membranes [193,194]. The lethal action of polymyxin is attributed to the cytoplasmic membrane damage as a result of the interaction of polymyxin with the acidic phospholipids [193]. Also the outer membrane binds polymyxin [194] and subsequently looses its permeability barrier function. A clear effect of the antibiotic on the structure of the outer membrane has been shown [ 195,196]. Initial binding to the outer membrane most likely is an interaction with LPS [142,177,197], presumably with its anionic KDO or phosphate groups [197].

EDTA removes about half the LPS of the cell [85], presumably by complexing divalent cations which are involved in LPS-LPS interactions. In our hands the procedure does not result in removal of significant amounts of outer membrane protein [198]. The presence of Tris ions enhances the removal of LPS from the cell by EDTA [199]. Interestingly, a direct interaction between the Tris ion and the LPS molecule has been reported [197].

Divalent cations have a high affinity for LPS [85,197,198,200]. Schindler and Osborn [197] have shown that there are two binding sites for Mg 2+ and Ca 2+ . The first one, of relatively low affinity, was attributed to pyrophosphoryl and /o r phos- phodiester groups of the KDO - lipid A region. The second site, of higher affinity, must probably be attributed to the branched KDO trisaccharide

unit [197], which therefore also is supposed to be required for assembly or maintenance of the normal structural organization of the outer membrane [197]. These affinity sites for divalent cations on the LPS molecule could play an important role in the assembly or maintenance of the molecular organization of the outer membrane by bringing about LPS-protein an d /o r LPS-LPS interactions.

IIID. Enterobacterial common antigen (ECA)

The enterobacterial common antigen or Kunin antigen is shared by most Enterobacteriaceae [201] and is an often forgotten constituent of their outer membranes [202]. ECA represents a polymer of N-acetyl-D-glucosamine and D-mannosaminuronic acid, partly esterified by palmitic acid [203]. Its chemistry and biology have been extensively re- viewed [201,204]. In addition to the free form a few strains contain ECA also in the immunogenic form in which it is associated with lipopolysaccharide by an interaction to the core-lipid A part, similar to the way the O-antigen is linked.

IIIE. Proteins

lllE-1. Introductory remarks As the outer membrane is very poor in en-

zymatic activities [76], the identification of the protein component is mainly dependent on separation of proteins in bands using SDS-polyacrylamide gel electrophoresis. Compared with the present systems which give a high resolution [205- 209] the systems used earlier were rather poor. Also the introduction of slab gels [91,210] has largely contributed to the resolution of the outer membrane proteins. Two-dimensional gels, which have given new opportunities for studying complex mixtures of soluble proteins [211] have been applied to outer membranes in only a few cases [212-214] and turned out to give rise to a large number of artefacts due to strong interactions between outer membrane constituents (see Ref. 213). The molecular weight of a protein can be estimated by SDS-polyacrylamide gel electrophoresis. As a proven or supposed single amino acid substitution can result in a drastically altered electrophoretic mobility [215-219], the apparent molecular weight value obtained should be interpreted with care.

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66

Initially, outer membrane protein patterns obtained in various laboratories differed drastically, partly because different solubilization temperatures were used during the preparation of the sample, and partly because different gel systems were applied. Since some interactions between outer membrane constituents are extremely strong, temperatures above 70°C are required for complete solubilization of the outer membrane proteins of E. coli and S. typhimurium and the standard procedure used in these cases now consists of boiling the sample in 2% SDS for a few minutes. In the cases of H. influenzae (Van Alphen, L., unpublished data) and Pseudomonas aeruginosa [220] complete unfolding of all outer membrane protein chains even requires boiling for 10 and 30 rain, respectively. Incomplete unfolding of a polypeptide chain can result in binding of insufficient amounts of SDS. (Usually as much as 1.4 g of SDS is bound per g protein!). Therefore incomplete solubilization can result in an altered electrophoretic mobility or even in the absence of a protein from the running gel, a phenomenon which is often accompanied by the accumulation of protein on top of, or in, the stacking gel. The altered electrophoretic mobility of a number of incom- pletely unfolded proteins has resulted in the term heat-modifiable proteins which can often be found in the earlier literature.

The relatively poor resolution of the earlier gel systems has introduced the notion that one protein was supposed to account for 70% of the total outer membrane protein. Subsequently this 'major outer membrane protein'[86] of E. coil K-12 was resolved into two [221,222] and later on into four [205,207] proteins (Table I), each of which is still designated as a 'major outer membrane protein' although the relative abundance of one or two of these proteins usually is very moderate [205,232]. Moreover, the term is misleading as the lipoprotein, which is by far the most abundant protein of the cell, does not belong to the 'major outer membrane proteins'! Finally, the term 'major ' is often relative as in several cases the growth conditions influence the amount of a given protein. The observation that the amount of major outer membrane protein per unit of outer membrane surface area is constant under various growth conditions [233] is important. Firstly, it shows that a regulation mecha-

nism is operative for major proteins and secondly, it provides us with a practical standard for cell surface area, which as such is tedious to measure. It means for example that our previous data on the amounts of individual proteins relative to the total amount of outer membrane protein [95] can be extended to the amount of the individual proteins per unit of cell surface (see later on).

The use of different SDS-polyacrylamide gel electrophoresis systems in different laboratories made a comparison of the early data difficult, resulting in several different nomenclature systems (Table I). The use of improved gel systems, the availability of mutants lacking one or more proteins and the availability of pure preparations of several proteins are the main factors responsible for resolution of this confusion. In the case of E. coli it was finally agreed upon by most groups to use a uniform nomenclature system in which the protein was named after its structural gene [4] (Table I). Thus, a protein previously known as If*, d, 3a, B, TolG protein or O10 is now called OmpA protein as gene ompA is the structural gene for this protein. (The genetic abbreviation omp stands for outer membrane protein). Although a generally accepted nomenclature system certainly is an improvement, its coupling to the genetic code is a severe restriction. For example, as the structural gene of the protein known as a, 3b or O l l (see Table I) is not known with certainty, the protein cannot be properly named despite the fact that its purification has been described alreay in 1974 [230a]. Also, the system cannot easily be used for other strains than E. coli K-12. For example, certain E. coli strains have as many as five proteins which are immunologically related with OmpC protein and OmpF protein of E. coli K-12. It would probably take years to find out whether one of the proteins of the wild strain is indeed coded by the presumed ompF gene of this strain, and if so, which one.

Several outer membrane proteins are designated as 'peptidoglycan associated' proteins. This term is correct as long as it is used in the operational sense, namely as proteins which under certain conditions (e.g. in 2% SDS at 60°C) are the only proteins which remain strongly but non-covalently bound to the peptidoglycan fraction. The question of whether this is also the case in vivo will be

67

handled in subsection IVF-5. A comparison of cytoplasmic and outer mem-

brane preparations by SDS-polyacrylamide gel electrophoresis shows relatively few, but often heavy, bands in the case of the outer membrane. It should be stressed that this result not necessarily means that there are only a few outer membrane proteins as some proteins might be present in amounts below the level of detection, e.g. phospholipase A1 and the BtuB protein can hardly or not at all be detected as bands in crude outer membranes, whereas other proteins may be present in the outer membrane in detectible amounts only under certain growth conditions (e.g. inducible proteins from cells grown under conditions which repress their synthesis, see subsection IIIE, 5, 6 and 7).

Genetics has played an important role in the fast increase of knowledge on E. coli outer membrane proteins. As several proteins are used as a receptor by bacteriophages, mutants resistant to such a phage often lack the receptor protein. Com- parison of properties of mutant and parent has often indicated the function(s) of the protein (see also subsection IVA for interpretation). Mutant cells have been used as the starting material for the purification of proteins as mutants can be constructed such that an otherwise persistent impurity is lacking or such that they overproduce the required protein. Mutants are indispensible for studies on the regulation of the synthesis of specific proteins. A recent genetic development is the con- struction of operon and gene fusion strains which can be used as powerful tools for studies on the regulation of transcription [234,235] and on the subcellular localization of proteins [11,14,16], respectively. The availability of mutants is usually also useful for molecular cloning of the corresponding wild-type gene. Cloned DNA can be used for detailed studies on regulation and expression of the gene product. Moreover, cloning not only enables the investigator to determine the nucleotide sequence of a structural gene and thus to predict the primary structure of the protein but it also provides the possibility to determine the signal sequence and to study the regulatory regions and other aspects of the operon structure.

A single mutation sometimes results in decreased amounts of several outer membrane pro-

teins. For example, mutations causing a heptose- less LPS result in strongly decreased amounts of OmpF protein [94], PhoE protein [236], protein III [93] and Lamb protein [237] in the outer membrane. Using an ompF-lacZ operon fusion strain it could be shown that in the case of OmpF protein poor expression was not the result of poor transcription (Tommassen, J., Overduin, P. and Lugtenberg, B., unpublished data). Therefore it is likely that the defects are caused by a poor assembly or translocation of the proteins.

Another indication for a structurally common site on various exported proteins comes from work with mutants containing a defective perA gene. These cells have decreased amounts of alkaline phosphatase, of several other periplasmic proteins [238], and of the outer membrane proteins OmpF protein [238], LamB protein [239], PhoE protein (Tommassen, J., Beusmans, J. and Lugtenberg, B., unpublished data), protein a, Cir protein, FepA protein and 83 kDa protein [240].

The following sections will focus on the structural and functional characteristics of outer membrane proteins. All predominant outer membrane proteins have been purified and in many cases biological activity depends on the presence of LPS. Outer membrane proteins are usually rich in fl- structure [241]. It appears that certain proteins play a role in the stabilization of the structure of the outer membrane and in anchoring this membrane to the peptidoglycan layer. Other proteins facilitate the permeation of nutrients through the outer membrane. Some proteins seem to be produced constitutively whereas the synthesis of others is dependent on the growth conditions.

IllE-2. Enzymes in the outer membrane The outer membrane is poor in enzymatic activ-

ities [76,242,243]. Phospholipase A1, M r 28000 [244,245], is the first enzyme detected in the outer membrane [76,242]. Later on, lysophospholipase, lysophosphatidic acid phosphatase and UDP-glucose hydrolase were found in this membrane [82]. The first example of an enzyme activity which is clearly present in both membranes comes from Vos et al. [246] who reported that the specific activity of monoacylglycerophosphoethanolamine acylase, which does not require ATP or coenzyme A for activity, is the same in cytoplasmic and outer

68

membrane preparations of E. co#. This location in both membranes makes sense as reacylation of lysophospholipids formed by the action of endogenous or exogenous phospholipase A provides the organism with the potential of biochemically inex- pensive repair and modification of the envelope phospholipids. Moreover, major phospholipids hydrolyzed in the outer membrane can be resynthesized in the same location, without need for transport of the products of hydrolysis to the lipid biosynthetic apparatus in the cytoplasmic membrane [246].

The outer membrane was found to contain proteolytic activities able to convert the precursor of alkaline phosphatase to the mature form [247], to modify the ferric enterobactin receptor [248], to hydrolyze casein [249], to cleave bacteriophage M13 precoat protein to coat protein [250], to solubilize nitrate reductase [251], to cleave colicin Ia [252] and to activate serum plasminogen to active protease plasmin [253]. Presently it is not known how many enzymes are responsible for these activities but two of them have recently been purified.

The casein-hydrolyzing enzyme, designated as protease IV, has been purified by Regnier [254,255 ], It is an endoproteolytic enzyme with a M r of 23 500 which is mainly localized in the outer membrane. It can be solubilized by deoxycholate or SDS, it is resistant to thermal denaturation and is inhibited by EDTA, by various protease inhibitors and by inhibitors of processing enzymes [254,255]. A localization on the inner side of the outer membrane has been suggested [254].

The second enzyme which has been purified converts M 13 precoat protein to coat protein [256]. This processing enzyme has been designated as leader peptidase or signal peptidase. It has an apparent molecular weight of 39 000 and does not require cofactors for activity [256]. It is the second enzyme which has been found in equal abundance in the cytoplasmic and outer membranes of E. coli [257], The enzyme also cleaves precursor forms of two periplasmic amino acid binding proteins and of the Lamb protein [257]. A 4-6-fold overproduction of leader peptidase was observed in a strain bearing plasmid pLC7-47, presumably due to the presence of several copies of the structural gene on the plasmid. This overproduction was not

accompanied by an altered cytoplasmic or outer membrane protein pattern [258]. Currently the precise role of leader peptidase in the assembly of M13 precoat is elegantly being studied in vitro [259,260].

Experiments of Wolf-Watz and Normark [261] suggest that a peptidoglycan hydrolytic enzyme, N-acetylmuramyl-L-alanine amidase, is loosely associated with the outer membrane.

It has recently been reported that Serratia

marcescens, a Gram-negative bacterium which ex- cretes lipase, protease and nuclease activities [120,262], can contain substantial quantities of these enzymes in its outer membrane, presumably an intermediate location before excretion occurs [263]. Even more striking is the observation that in the same organism about 80% of the fl-lactamase activity is found in the outer membrane. Finally, outer membranes of Neisseria meningitidis contain tetramethylphenylenediamine oxidase activity [264].

l l lE -3 . Lipoproteins Braun and co-workers [265] were the first inves-

tigators who purified an outer membrane protein. This lipoprotein from E. coli ( M r 7200) contains 58 amino acid residues, and is covalently bound to the carboxylgroup of every tenth to twelfth diaminopimelic acid residue of the peptidoglycan layer through the e-NH 2 group of its C-terminal lysine residue (for a review see Ref. 266). This covalently bound form of the lipoprotein is present in approx. 2.4-105 copies per cell [266]. It was subsequently discovered that twice as many copies of exactly the same lipoprotein molecule exist in the free form i.e. not covalently attached to the peptidoglycan [266a]. The lipoprotein is by far the most abundant protein of the cell. Both forms of the lipoprotein have been purified and sequenced [267,268] and the free form has been crystallized [268a,268b]. The amino acids histidine, tryptophan, glycine, proline and phenylalanine are lacking. As the amino acid sequence of the lipoprotein is highly repetitive it was speculated that the lipoprotein is evolved from a 15 amino acids long peptide by gene multiplication and subsequent mutations [266]. The N-terminal cysteine residue is substituted both with a diglyceride moiety in a thioether linkage as well as with an amide-linked fatty acid.

69

The isolation of a lpp mutant lacking the structural gene for both forms of the lipoprotein located at rain 36.3, shows that the lipoprotein is not esential for survival of the cell. However, a lipoprotein deletion mutant has severe defects like increased production of outer membrane vesicles, increased sensitivity to EDTA and leakage of periplasmic enzymes, suggesting a role in the stabilization of the outer membrane [269,270]. When in addition to the lipoprotein also the OmpA protein is missing, cells are unable to grow in the rod form and require high concentrations of Mg 2÷ or Ca 2÷ for growth [271]. Moreover abundant blebbing was observed in the double mutant and the peptidoglycan layer was no longer connected with the outer membrane [271] suggesting a role of the proteins in the determination or maintenance of the rod shape, in stabilization of the outer membrane structure, and in anchoring the outer membrane to the peptidoglycan. Additional evidence for the latter function comes from recent experiments of Wensink and Witholt [272] who showed that outer membrane vesicles released by growing E. coli cells contain only a small amount of free lipoprotein, hardly any bound lipoprotein and reduced amounts of OmpA protein. The vesicles also contained reduced amounts of protein V, a protein assumed to be identical to a newly discovered lipoprotein [272].

The lpp genes of E. coli K-12 [273], Serratia marcescens [274] and Erwinia amylovora [275] have been cloned in lambda phage vectors and their nucleotide sequences have been determined. Al- though E. coli is only distantly related to the other two species, comparison of these sequences suggests that the structure of the lipoprotein has been highly conserved [275]. Attempts to clone the E. coli lpp gene into the high copy number vector pBR322 have not been successful, presumably due to lethal overproduction of the lipoprotein [276]. In contrast, the lpp gene of S. marcescens could be cloned into this vector and the resulting plasmid restored the wild-type phenotype of an E. coli lpp mutant. This Serratia lipoprotein was exclusively found in the outer membrane. The effect of multiple copies of the structural gene resulted in a 3-fold overproduction of the free form of the lipoprotein compared with E. coli lpp + cells, whereas no overproduction of the bound form

lipoprotein occurred [276]. A similar gene dosage effect for the free form, but not for the bound form was observed in E. coli cells carrying an F' factor containing the Ipp gene [277].

With respect to the 3-dimensional structure it has been determined that the lipoprotein, in contrast to many other outer membrane proteins, is very rich in a-helix [278,279] which is consistent with Inouye's model in which all hydrophobic amino acid residues are regularly arranged in an alternating 3 to 4 pattern of repeating hydrophobic residues. As 3.6 residues make up one regular right-handed a-helical turn, all the hydrophobic residues can be alligned as two series on one face of the helical rod [266,280].

The structure of the lipoprotein has been extremely well conserved in Enterobacteriaceae [274,275,281-284]. Lipoproteins have also been found in Aeromonas salmonicida (Evenberg, D., Overbeeke, N. and Lugtenberg, B., unpublished data), Pseudomonas aeruginosa [285] and in Rhodopseudomonas spheroides [286]. The presence of lipoprotein has not been established in other bacteria. For example, it could not be identified in N. gonorrhoeae, grown at pH = 7, which might explain the strong blebbing of the outer membrane which seems only loosely attached to the peptidoglycan layer [287].

Another class of lipoproteins, peptidoglycan-associated lipoproteins or PAL proteins, which occur closely but non-covalently associated with peptidoglycan, has recently been found in various Gram-negative bacteria like P. mirabilis [288], Ps. aeruginosa [288,289] and E. coli [290]. They are not immunologically cross-reactive with Braun's lipoprotein [291]. PAL of P. mirabilis has an M r of 18 000 and its N-terminus, including the lipid part, in common with Braun's lipoprotein [288]. Very recently several new lipoproteins, which are immunologically different from both Braun's lipoprotein and PAL, have been discovered in E. coli [290]. From these new lipoproteins four were located in the outer membrane and two in the cytoplasmic membrane, whereas the localization of one species is uncertain. Thus the total number of biochemically different lipoprotein species in E. coli is at least nine [290]. Interestingly, Inouye and co-workers [292] come to a similar conclusion on the basis of DNA hybridization experiments.

70

llIE-4. OmpA protein OmpA protein is present in about 105 copies

per cell. It is heat-modifiable in that its apparent molecular weight on polyacrylamide gels is higher in the heat-modified form (35000) than in the non-denatured form (28000) [221,224,241,294]. In two-dimensional gels the protein appears in at least 12 spots, most of which are caused by artefacts [2131.

A procedure has been described for the purification of four major proteins from one batch of cells of E. coli strain B / r [294]. As this procedure applied to E. coli K-12 resulted in large losses of OmpA protein in our hands, we have modified the method in order to purify the E. coli K-12 protein [293]. The OmpA protein has a high B-structure content [241]. Purified OmpA protein is not partic- ularly hydrophobic [293,295,296]. Although it has been reported previously that OmpA protein contains glucosamine [297], it was claimed later on that non-protein substituents were not detected [295,296]. The complete amino acid sequence of OmpA protein of E. coli K-12 has been determined [296]. It consists of 325 residues resulting in a M r of 35 159 [296]. Computer analyses did neither reveal internal homology nor homology with OmpF protein [296]. The high electrophoretic mobility of the non-heat modified form is ascribed to the high content of B-structure and excessive binding of SDS in the absence of heating in SDS [296,298]. Of the lysine residues of OmpA protein 6-24% are present as allysine (a-amino adipic acid semialdehyde) as a result of an enzymatic [299] post-transcriptional modification process [300]. It has been suggested that a possible function could be to crosslink the protein to diaminopimelic acid residues of the peptidoglycan layer since some OmpA protein is covalently linked to this layer in stationary phase cells [300].

A protein cross-reactive with OmpA protein was detected in all strains of E. coli [301,302], of other Enterobacteriaceae [302,304] and also in Aeromonas salmonicida (Evenberg, D. and Lugten- berg, B., unpublished data) and Haemophilus influenzae (Van Alphen, L. et al., unpublished data).

OmpA protein deficient mutants of E. coli can be obtained by selection for resistance to phages K3 [305] or TuII* [306] or by isolating mutants tolerant to bacteriocin JF246 (colicin L) [225]. The

lack of OmpA protein is compensated for by increased amounts of pore proteins [293,306] and phospholipid [127] and possibly also of LPS [127]. The mutants do not adsorb the mentioned phages and are defective in F-pilus mediated conjugation [6,305,307]. The suggested receptor or receptor-like activities could be mimicked with purified OmpA protein provided that it was complexed with LPS [293,308]. The role of OmpA protein in conjugation was confirmed by independent means by Havekes and Hoekstra who selected mutants defective in the acceptor function in F-pilus mediated conjugation and subsequently showed that the mutation was localized at the ompA locus [307]. Its function in conjugation most tion of mating aggregates Braun's lipoprotein OmpA involved in maintaining both ity of the outer membrane as of the cell [271].

likely is the stabiliza- [6]. Together with

protein is somehow the structural integr- well as the rod shape

The structural gene ompA, located at rain 21.5 [216] of the E. coil K-12 chromosome, has been cloned [309,310]. Expression of a few copies of the gene results in a 2-fold overproduction of the protein [309] whereas a further increase in overproduction is lethal for the cell [309,311]. The overproduction of OmpA protein as a result of cloning causes decreased levels of OmpF protein, OmpC protein, LamB protein and lipoprotein in the cell envelope [309]. It is interesting to note that by genetic manipulation of the cloned material a 30 000 dalton fragment, which lacks the 96 [310] or 98 [311,312] -COOH terminal residues, is incorporated in the outer membrane. As the presence of the fragment makes the cells sensitive to the OmpA protein specific phages it can be concluded that the carboxyterminal third of the protein is not required for incorporation in the outer membrane [310,3111.

Although purified OmpA protein is completely degraded by proteolytic enzymes, trypsin [313] and pronase [312] leave N-terminal parts of M r 24000 and 19000, respectively, intact as long as the protein is embedded in the membrane. This large fragment is still able to act as a phage receptor whereas it most likely also still functions in F-pilus mediated conjugation [312,313]. Results with proteolytic enzymes [313] and with shortened gene fragments [312] indicate that the OmpA pro-

tein consists of two domains. The N-terminal moiety from residues 1 to 180 represents the membrane domain whereas the remaining 55 residues are proposed to be located in the periplasmic space [312].

Several investigators have studied the expression of ompA genes in E. coli and other bacteria. E. coli K-12 strains harbouring episome F'106 containing the ompA gene, do not produce more of this protein. Transfer of this episome to S. typhimurium and P. mirabilis resulted in incorporation of the E. coli OmpA protein in the outer membranes of the recipients [314]. Similarly, cloned ornpA genes of Shigella dysenteriae, Enterobacter aerogenes and Serratia marcescens are fully expressed in the outer membrane of E. coli K-12 [315 ]. These results are hard to reconcile with observations from Schaitman's laboratory which indicate that ompA genes from a certain E. coli strain are not expressed in other E. coli strains unless the cloned ompA gene has been mutagenized in such a way that the affinity of the OmpA mutant protein for the LPS of the recipient strain has increased [304].

By comparing nucleotide sequences of cloned ompA genes of various bacteria, Henning, Cole and their co-workers are trying to correlate differences in structure with differences in biological activities [312,316].

IIIE-5. The family of peptidoglycan-associated general diffusion pore proteins

I11E-5a. Introduction. When whole cells or cell envelopes of E. coli strain B E are incubated at 60°C in 2% SDS and subsequently centrifuged, the pellet consists of peptidoglycan with the lipoprotein covalently bound to it and another protein with a M r of 36 500 noncovalently bound to it. The latter protein can be removed from the peptidoglycan-lipoprotein complex by incubation in 2% SDS at a temperature of 70°C or higher [230] or by incubation at 37°C in SDS containing 0.5 M NaC1 [317]. Heating results in changing of the tertiary structure from B-sheet to a-helix thereby irreversibly denaturating the protein. Release of the protein by high salt leaves the native confor- mation intact. As the peptidoglycan-lipoprotein complex of E. coli strain B E with, but not without, the non-covalently 'peptidoglycan-associated' pro-

71

tein morphologically appears to contain a hexagonal lattice structure, the protein was designated as 'matrix protein' [230]. Later on the latter term has often been used for similar proteins or even for mixtures of proteins. However, as the experiments leading to the designation 'matrix protein' have only been described in the case mentioned above [230], the use of the term 'matrix protein' for these other proteins should be discouraged. We prefer the term' peptidoglycan-associated protein' as long as it is used in the operational sense (see subsection IIIE-1).

E. coli strain K-12 contains two peptidoglycan- associated proteins [95,228,318], known as OmpC protein and OmpF protein. They are immunologically related with each other as well as with PhoE protein [319], an inducible pore protein in this strain [320]. A survey of 45 hospital isolates of E. coli has shown that the number of peptidoglycan- associated proteins per strain in the M r range between 30 000 and 42 000 can vary between one and four, whereas the electrophoretic pattern of these proteins was OK serotype-specific [301a,b]. The denatured form of each of these peptidoglycan-associated proteins reacted with at least one of the antisera raised against highly purified de- naturated OmpC protein, OmpF protein or PhoE protein. As none of the other E. coli cell envelope proteins reacted, these results suggest that all these pore proteins are derived from a common ancestral gene and they show that the structures of these genes have been conserved very well during evolu- tion [301]. Peptidoglycan-associated proteins of Enterobacteriaceae are very common in nature. Three of them have been detected in S. typhimurium [90,321] and a large survey has shown that they are present in all Enterobacteriaceae tested [322-324] and in Ps. aeruginosa [325] and that also these proteins are cross-reactive with the E. coli proteins [301-303,326]. The family of peptidoglycan-associated proteins is even larger if one takes into account that not all proteins are constitutively present but some are induced under certain growth conditions only [320,327] or are coded for by a (pro)phage [328,329] or by a plasmid [330-332]. It should be noted that the temperature of 60°C is arbitrary. If lower temperatures are used additional proteins are found to be associated with peptidoglycan in E. coli K-12 [333] and Proteus vulgaris [322].

72

TABLE il

CHARACTERISTICS OF SOME E. COL1 AND S. TYPHIMURIUM PEPT1DOGLYCAN-ASSOCIATED GENERAL DIFFU- SIONS PORE PROTEINS a

Protein species Mr Number of copies/cell (Part of) receptor for phage/bacteriocin

Structural gene Purification (Refs.) lsoelectric point (pH) Refs. to amino acid composition and sequence Oligomeric form Pore activity (Refs.) Pore diameter (nm) Further characteristics

OmpF protein (E. coil strains K-12 and B) 37 205 [376] Up to 105 [230] Tula [381], T2 [382], TP1 [383], K20 b, TP2 [384], TP5 [384], cola [384a]

ompF, min 20.7 [392-396] 230, 322, 396 5.9-6.2 [230]

227, 230, 374, 376, 396 Trimer [356,397] 339, 341,342, 343, 344, 358 1.4 [358] Gene ompF has been cloned [395,402]; it hybridizes with the phoE gene [395]; synthesis of OmpF protein is repressed by high osmolarity [403,404]; synthesis positively controlled by cAMP? [405]; 70% amino acid sequence homology with phoE protein [380]

OmpC protein (E. coli K-12) 36000 [205] Up to l0 s Tulb [381,385], T4 c [386,3871, Mel [3881, PA-2 [329], 434 [382], SS1 [389], TP2, TP5, TP6 [384] ompC, min 47.1 [217,231,388,396] 322, 396 n.d. 0

227, 374, 396 Trimer [397,398] 342, 343, 352, 358 1.3 [3581 For effect of osmolarity see under OmpF protein. Smaller effective diameter than OmpC pore [342,343,409]

a Association with peptidoglycan is meant in the operational sense only (see subsections lllE-5a and IVF-5). The group of proteins listed in this table has in common an antigenic relation with OmpF protein and/or OmpC protein of E. coli K-12 [319].

h Manning, P. and Reeves, P,, personal communication.

l l IE-5b. Function of general pore proteins. E.

coli has many high affinity t ransport systems in its cytoplasmic membrane with K m values around 1 /~M for most solutes [334]. If such a t ransport system also has a reasonably high Vma X value the outer membrane will form a serious t ransport barrier at low solute concentra t ions unless it contains an extremely high number of channels. Nikaido and Nakae and their co-workers have developed the concept of hydrophilic or water-filled pores in the outer membranes of Enterobacteriaceae in order to explain both the outer membrane ' s impermeability for bile salts and its extremely good permea-

bility for nut r ients and other solutes with a M r up to approx. 600 [335-337]. Although most func-

t ions of bacterial const i tuents have been unmasked with the help of mutants , the function of peptidoglycan-associated proteins was discovered by purif icat ion of the outer membrane componen t responsible for the generation of aqueous pores in phospho l ip id -LPS l iposomes through which galactose, g lucosamine , g lucose - l -phospha t e , leucine, glutamic acid, lysine, t ryptophan, uridine, UMP, G D P and poly(ethylene glycol) (Mr approx. 600) could pass bu t which were impermeable for poly(ethylene glycol) ( M r 1540). In the case of S.

73

TABLE 11 (continued)

CHARACTERISTICS OF SOME E. COLI AND S. T Y P H I M U R I U M PEPTIDOGLYCAN-ASSOCIATED GENERAL DIFFU- SIONS PORE PROTEINS a

PhoE protein (E. coli K-12) OmpF protein [4] OmpC protein [4] OmpD protein [4] (35 kDa protein) (36 kDa protein) (34 kDa protein) ( S. typhirnurium ) ( S. typhimurium ) ( S. typhimurium )

36782 [380] 39300 [321] 39800 [321] 38000 [321] Up to 105 Up to 105 Up to 105 Up to 105

TC23, TC45 [226,390] PH42, PH105, PH221 [391] PH31, PH42, PH51 [391] phoE, min 5.9 [218] ompF, min 21 [4] ompC, min 46 [41 ompD, min 28 [4] 226,236 321 321 321 n.d. 4.77 [321] 4.78 [3211 4.85 [3211

227,236,374,380,396 n.d. 236,343,345,358,400 1.2 [358] PhoE protein synthesis is derepressed by Pi- limitation [320]; PhoE protein pore is extremely efficient for Pi and organic P [345] due to a recognition site [400]; 70% amino acid sequence homology with OmpF protein [380]; gene phoE has been cloned [406]; overproduction of PhoE protein is lethal [406]; phoE gene hybridizes with ompF gene [395]

321 321 321 Trimer [399] Trimer [357,399] Trimer [357,399] 340,401 340,401 340,401 1.4 [4011 1.4 [4011 1.4 [4011 Synthesis repressed by salt [34]

c Whereas E. coli B LPS alone is sufficient for phage inactivation [379] a complex of LPS and OmpC protein is required in the case of E. coli K-12.

d n.d., not determined.

typhimurium the active fraction contained the three peptidoglycan-associated proteins 34, 35 and 36 kDa whereas fractions containing the 33 kDa protein (comparable with the OmpA protein of E. coli) or the free form of the lipoprotein were inactive [338]. Subsequently Nakae could show that the only peptidoglycan-associated protein of E. coli strain B had the same general diffusion pore activity. He therefore proposed the term 'porin' for such proteins [339]. Subsequent in vivo work using mutants lacking one or more peptidoglycan-associated proteins confirmed the concept of the pore function for these proteins

[340-344] and enabled one to measure rates of penetration which at that time could not be measured with the liposome system. Such in vivo studies using carefully designed mutants provided the first indication for functional differences between the E. coli K-12 pore proteins OmpC protein, OmpF protein and PhoE protein [342,343,345,346]. In fact, pore protein deficient mutants have an increased K m and an unaltered Vma ~ for the uptake of solutes [9,341,345-348]. A large improvement was the development of rate measurements in vitro by the use of black lipid films [349,350], by application [351,352] of the liposome swelling

74

method [353,354] and by the incorporation of a specific solute-converting enzyme into liposomes [355]. The biologically active form of the porin turned out to be a trimer [356,357].

The most important conclusions from the work on the functioning of pores can be summarized as follows.

(i) The simplest interpretation of a pore is a nonspecific molecular-sieving channel through which hydrophilic solutes diffuse. The diffusion rate is determined by the difference in concentration at the two sides of the membrane. The temperature dependence of the rate of penetration is low [337,338].

(ii) Solutes which are too bulky cannot diffuse through a pore. In Enterobacteriaceae the size limit for oligosaccharides is 600-700 daltons which cor- responds with a pore diameter of 1 nm [9]. From conductivity measurements through black lipid films, and assuming that the pore is a cylinder of 7.5 nm length, the proposed thickness of the outer membrane, pore diameters of 0.9-1.4 nm were calculated [9,350,358].

(iii) The diffusion rate of solutes through pores is influenced by factors like size, charge and hydrophobicity, resulting in large differences in permeability coefficients among solutes [9,10,359].

Using the procedure developed by Zimmerman and Rosselet [360], it is possible to measure rates of permeation of ,~-lactamase sensitive/3-1actams through pores in the outer membrane. By applying this procedure on a series of B-lactams with different hydrophobicity, Nikaido showed that the permeability coefficient increases with decreasing par- tition coefficient in an isobutanol-aqueous buffer system [9]. Cephaloridine behaved anomalously as it penetrated approx. 6-times faster than predicted [9]. The results certainly show the importance of hydrophilicity for penetration through pores. Ap- parent anomalous behaviour must be due to our too simple picture of pores as just holes. In this respect it is important to note that it has recently been shown that the channels formed by LamB protein and PhoE protein have a preference for certain substrates due to the presence of recognition or binding sites, presumably at the entrance of the pore (see subsections IIIE-6b and IIIE-5dii, respectively).

Enterobacteriaceae are relatively resistant to /~-

lactam antibiotics whereas many other Gram- negative bacteria are sensitive. In the latter case it is assumed (see section IV) that the antibiotics mainly use the hydrophobic pathway, i.e. they diffuse through the phospholipid bilayer [9,361]. Nikaido [361] has proposed that the hydrophobic pathway does not exist in the outer membrane of Enterobacteriaceae and that in this case fi-lactams are dependent on pores for permeation [361]. With respect to chemotherapeutic use of fl-lactam antibiotics against Enterobacteriaceae it is interesting to note that the permeability coefficient of cepha- cetrile and cephaloridine is only 3 4-fold lower than that of lactose. Thus, Nikaido concluded that the goal of making very hydrophilic, and therefore rapidly pore-penetrating, /3-1actams has already been achieved for these compounds [10].

(iv) The incorporation of pore proteins into planar lipid bilayer membranes leads to an increase of the membrane conductance of many orders of magnitude [349,350]. At lower protein concentrations the conductance increases in a stepwise fashion. These findings are consistent with the assumption that the protein forms large aqueous channels in the membrane. The formation of channels is induced irreversibly by voltage. The channels exist in either an open or a closed state, which are in equilibrium with each other. Interest- ingly, cooperativity among channels was observed and the smallest inducible unit presumably corresponding to a trimer, consists of three channels [350,362]. This result, together with structural studies indicating that a functional pore consists of three monomers [356] strongly suggests that each monomer can form a pore and that opening of the three channels within a trimer is a highly cooperative phenomenon.

LPS is required for channel activity [362], a conclusion not unexpected as a requirement for LPS has been shown for practically all biological activities of pore proteins (see subsection IVB-3). The assumption that LPS is not required in black lipid films [349] must be due to the misunderstanding that the used isolation procedure yields completely LPS-free pore protein.

(v) In strains containing more than one pore protein species the removal of all but one of these species (by mutation or by the choice of the right growth condition) leaves the cell with a perfectly

75

functional pore [342,343,363,364] showing that also in these cases one pore protein species is sufficient.

(vi) The term porin has been introduced to indicate general diffusion channels [339]. However, it is known now that some pores, in addition to having general pore properties, exert some preference for solutes with respect to the rate of permeation. For example, evidence has been presented that the LamB protein pore (see subsection IlIE- 6b) and the PhoE protein pore (see subsection IIIE-5dii) have recognition sites for certain solutes, resulting in a faster permeation of these solutes. Next to a built-in recognition site another mechanism has been proposed to explain pores which exert a preference. Lo and Bewick [365-368] have recently provided strong evidence that an inducible dicarboxylate binding protein is located at the outside of the outer membrane. The observation that succinate uptake at a concentration of 1.75 /~M is virtually abolished in a mutant lacking both OmpC protein and OmpF protein led the authors to suggest that the binding protein is physically connected with the outside of the pore and serves as a substrate recognition component [365]. Work to test this interesting possibility should be pursued.

(vii) A study of the permeability properties of the outer membrane of Pseudomonas sp. is particu- larly interesting as these organisms are very resistant towards many antibiotics but are on the other hand able to utilize a large variety of nutrients. Although it was initially thought that the pores of Ps. aeruginosa had the same exclusion limit as those of S. typhimurium and E. coli [337], later experiments have shown that the exclusion limit is much larger, approx. 6000 daltons [115, 369], which probably is advantageous for the organism in that it permits the entry of small micelles and hydrophobic solutes and of large peptides into the periplasmic space [115].

Benz and Hancock have extensively studied the behaviour of purified pore protein F [220] in black lipid films [358]. Like pore proteins of Enterob- acteriaceae, protein F increases the membrane conductance of artificial lipid bilayers by many orders of magnitude, but protein F differs from the former pore proteins in two important respects. Firstly, it causes a relatively high conductance, from which a channel diameter of 2.2 nm can be calculated.

Secondly, the pore activity per unit weight of purified pore protein is about 100-fold lower. Al- though artefacts cannot be ruled out, the authors prefer to explain the latter observation by assuming that relatively few pores are open at any given time, thus contributing to the antibiotic resistance of the organism [358]. Moreover, it has been suggested [9,358] that because of their large pore size, pore proteins of Pseudomonas sp. are good models for other pores in nature [9,358] (see for example Ref. 371).

Although it has been suggested that the increased resistance of Pseudomonas sp. to many antibiotics is the result of degrading enzymes or of reduced rates of transport through the cytoplasmic membrane [115,370], recent experiments using intact cells have shown that the rate of permeation of several solutes, including fl-lactams, through the outer membrane of Ps. aeruginosa is 100-fold lower than through the outer membrane of E. coli [378].

Pore proteins of Neisseria gonorrhoeae [372] and of other non-enteric bacteria [10] have been purified and studied in Nikaido's laboratory. They all had large channel diameters and it was therefore suggested that enteric bacteria might be rather exceptional in producing very narrow channels [101.

IIIC-5c. Purification and properties of general diffusion pore proteins. Most procedures used for the purification of pore proteins are based on the one developed by Rosenbusch [230], which is excellent for the preparation of chemically pure monomeric, denatured protein. However, if biologically active protein is required a high salt concentration should be used to remove these trimers from the peptidoglycan [317]. More recently deoxycholate [349] and fl- [362] or a-octylgluco- side [373] have given equally good or better results.

Denatured pore proteins are acidic polypeptides with isoelectric points varying from 4.8 [321] to 6.2 [230]. The amino acid composition of pore proteins is not hydrophobic [230,374]. The polarity coefficient of OmpF protein is as high as 45% [375]. As the purified trimer form is insoluble in water it is unclear whether the surface of the hydrophilic pore can accommodate the large number of polar amino acids and the possibility of ion-pair formation on the interior of the protein should certainly be examined [375]. In contrast to

76

the denatured monomer, the trimer form is rich in B-structure [230,357], binds relatively little SDS [230,356] and is resistant to proteases [230,338,357].

Like in OmpA protein (see subsection IIIE-4), some of the lysine residues of the E. coli K-12 pore proteins are present as allysine, although to a lesser extent (1-5%) [300]. These modified lysines were not detected during the determination of the complete amino acid sequence of the OmpF protein of E. coil B / r [376]. This polypeptide contains 340 amino acid residues resulting in a molecular weight of 37 205. The longest uninterrupted stretch of nonpolar residues consists of 11 amino acids. This fact as well as the percentages of polar amino acids in various regions of the molecule strongly suggest that the transmembrane region is not simply a single contiguous sequence of hydrophobic amino acids [376] as has been proposed for glyco- phorin [377].

lllE-Sd. Characteristics of individual general diffusion pore proteins. The characteristics of the best known general diffusion pore proteins of E. coli and S. typhimuriurn are summarized in Table II. Additional comments as well as more recent re- suits on these and other pore proteins are treated below.

i. OmpC protein and OrnpF protein (see also Table II). The early suggestion that OmpF protein and OmpC protein of E. coli K-12 were products of the same structural gene [228,231] together with the fact that these proteins are hardly or not at all separated in many gel systems [221,297] has led to the misunderstanding that E. coli K-12 contains one general pore protein. Even in more recent literature it occurs that these proteins are not differentiated (e.g. see Refs. 300, 407, 408). It is clear now that these constitutive pore proteins of E. coli K-12 are coded for by distinct but structurally related structural genes. The polypeptides are similar but differ in several properties [95,343,396]. The OmpF protein pore is larger than the OmpC protein pore [10,358,409].

The relative amounts of the two proteins are dependent on the composition of the growth medium, such that the sum of the amounts is almost constant [95]. Some evidence for a positive control of the synthesis of OmpF protein by cAMP has been published [405]. The osmolarity of the growth medium has a drastic effect [411] in that a

high osmolarity results in the disappearance of OmpF protein which is quantitatively compensated for by the synthesis of more OmpC protein [404]. Interestingly, solutes which cannot permeate the outer membrane exert this effect in substantially lower concentrations than those which pass this membrane [403]. The ompF gene of E. coli K-12 has recently been cloned [395,402]. Multiple copies of ornpF results in overproduction of OmpF protein at the expense of OmpC protein and PhoE protein [395]. Although transposon in- sertions have been obtained over the entire length of the gene, OmpF protein fragments have never been detected [395].

As both OmpA protein and OmpF protein interact with LPS, M o w a et al. [310] have compared the established and predicted sequences respectively and found several regions of similarity which are candidates for sites of interaction with LPS. It should be noted, however, that such an idea may be too simple as it could well be that different regions of LPS interact with the two proteins. For example it has been shown that OmpF protein is hardly present in the outer membrane of heptose- less mutants whereas the amount of OmpA protein is hardly affected by the mutation [94].

ii. PhoE protein (see also Table 11). This protein is induced in wild-type cells by phosphate limitation [320] as a component of a series of proteins [412] which the cell synthesizes in order to scavenge the last traces of phosphate or phosphate-containing components from the medium (for a review see Ref. 413). Consistent with this observation is the result that PhoE protein forms a more efficient pore for both inorganic and organic phosphate than OmpF protein [345], despite its smaller diameter [358]. This apparent discrepancy can be explained by assuming a recognition site for these solutes near the entrance of the pore. Strong evidence for this explanation has recently been presented [400]. Because of the structural similarities between PhoE protein and OmpF protein (see below) it thus is likely that PhoE protein has evolved from a general pore and, in order to play its role in phosphate uptake, has been provided with a weak binding site for certain solutes.

PhoE protein shares many properties with OmpC protein and OmpF protein (Table II). The proteins are immunologically related [319] and

77

recent work on the cloned genes ompF [395] and phoE [406] has shown that their DNA's can be hybridized over the entire length of the genes [395]. Such a homology at the DNA level could explain the large number of different pore proteins found in various strains of the same organism [301] as recombination between pore protein genes, in combination with mutation and transposition, could generate a large number of different but similar pores. The nucleotide sequence of the phoE gene has recently been established in our laboratory [380]. The predicted amino acid sequence was compared with the established sequence of the OmpF protein [376,414] and as many as approx. 70% of the residues, including several stretches of 15-20 amino acids, were identical [380]. Overpro- duction of PhoE protein, which is accompanied by decreased amounts of OmpF protein and Lamb protein, is lethal to the cells [406]. In contrast to the situation with OmpA protein (see subsection IIIE-4) expression of PhoE protein in the outer membrane cannot be detected if part of the gene, corresponding with the 50 carboxy terminal amino acid residues, is deleted [380].

Hancock et al. [416] have observed that phosphate limitation induces an oligomeric outer membrane protein, designated as P protein in Ps. aeruginosa. The observed in vitro pore properties of protein P show that it is highly specific for anions. Although protein P thus has important properties in common with PhoE protein of E. coli, it should be noted that pores formed by PhoE protein are significantly larger [416].

iii. Salmonella pore proteins (see also Table 1I). The Salmonella pores all have the same effective diameter [358] which is of the same size or slightly larger than those of the E. coli pores. The fact that the pores are equally effective makes the question why so many pores are needed even more intriguing. They have been compared with those of E. coli K-12 with respect to function, regulation of expression and, in the case of OmpC protein and OmpF protein, equivalence of the genetic loci (Nurminen, M. and M~ikela, P.H., cited in Ref. 4; Ref. 417). The results suggested that pairs of proteins corresponded (see Table II). The three proteins are immunologically [319] and chemically (Table II) related to E. coli K-12 pore proteins.

iv. Other general diffusion pore proteins. NmpC

protein, M r approx. 38 000, is produced by some colicin-sensitive revertants of porin-defective E. coli K-12 mutants. The mutation responsible for the reversion has been localized at min 12. By chemical and immunlogical criteria NmpC protein was reported to be similar to Lc protein, the former protein 2 [418]. In analogy to PhoE protein [320] it seems likely that NmpC protein is an inducible protein in wild-type cells.

Paakkanen et al. [419] examined the outer membrane protein patterns of 47 encapsulated and non-encapsulated E. coli strains. They observed that a protein band at the electrophoretic position of M r 40 000, designated as K protein, was present in all 33 encapsulated strains and absent in all but one of the 14 non-encapsulated strains. The authors therefore suggest that the protein is related to the capsule. The protein has been purified and resembles the OmpF protein of E. coli B/ r in its chemical properties as well as its apparent molecular weight [419]. The relation between the occurrence of the K protein and the capsule is very striking but the designation K (capsule) protein is prema- ture for two reasons. Firstly, it is based on the assumption that proteins with the same electrophoretic mobility have the same function. Sec- ondly, Paakkanen et al. have analysed the protein patterns on gradient gels in which the major proteins are present in a relatively narrow region of the gel. It would be interesting to see whether the relationship still holds when the patterns are analysed in a series of different gel systems. Using the homology between pore proteins [319,380,395] the structural gene can be cloned and be used to test the proposed relationship.

Lc protein [417], previously designated as protein 2, is coded by prophage PA2 [329,420]. When strains of E. coli K-12 are lysogenized with phage PA2 they replace the existing pore proteins OmpC protein (the receptor of the phage) and OmpF protein by the new pore protein [329], a form of lysogenic conversion at the protein level. The synthesis of Lc protein is catabolite repressible [329]. By chemical and immunological criteria Lc protein was reported to be similar to NmpC protein [418]. Recent work of the late Tom Gregg and co-workers [420] has shown that hybrid pore proteins, consisting of portions of Lc protein and OmpC protein, can be constructed. This type of

7~

approach can be very useful for studying structure-function relationships in pore proteins.

lyer [330,331] has observed that E. coli carrying an N plasmid produce an altered pore protein. We favour the explanation of Nikaido and Nakae [2] that this is caused by repression of the synthesis of the host pore protein(s) accompanied by synthesis of new plasmid-coded pore protein.

In Ps. aeruginosa a glucose-inducible pore protein has been described [421].

lIIE-6. Characteristics of E, coli pore proteins not antigenically related to the family of peptidoglycan-associated general diffusion pore proteins (Table 1II)

lllE-6a. Bacteriophage T6 receptor protein (Ta- ble 111). The receptor of phage T6 and colicin K, a protein of M r 26 000 [435,436], has been purified and its amino acid composition has been determined [436]. Biologically active T6 receptor in- variably contains LPS [436] and in vivo experiments suggest that core sugars play a role in a later step of the T6 infection process [422]. Dependence on LPS has been confirmed by in vitro experiments. The tsx gene is located at rain 9.2 and part of its has recently been cloned [438]. With respect to its function, the protein is involved in facilitating the diffusion of all nucleosides and deoxynuc- leosides except cytidine and deoxycytidine [422,423]. As the former solutes do not compete in the uptake process and as they do not inhibit T6 adsorption, it was concluded that the T6 receptor protein forms a pore to which the diffusing solute has only little, it any, binding affinity [422]. As the uptake system for nucleosides is catabolite repressible [435,439], the observation that the synthesis of the T6 receptor protein is also catabolic repressible [436] suggests that uptake of nucleosides is the real function of this protein. Experiments which suggest that the T6 receptor protein pore promotes diffusion of serine, glycine and phenylalanine (but not of glucose and arginine [440]) therefore are best explained as the acciden- tal use of this pore by these solutes.

lllE-6b. Bacteriophage lambda receptor protein (Table I11). The receptor of bacteriophage lambda, M r 47392 [424], is an outer membrane protein [425,441] which plays a role in the uptake of maltose and maltodextrins [442,443]. It is the

product of gene lamB, located at rain 91.0 of the E. coli genetic map, which is part of the well studied maltose regulon, which is used in model studies on the mechanisms of protein localization [444-447]. Addition of maltose to the growth medium derepresses the synthesis of the structural genes of this regulon. In addition to lambda [425] several other phages use the protein as their receptor (see Table Ill).

The lambda receptor protein is involved in maltose uptake as can be illustrated at low (approx. 1 /~M), but not at high (approx. l mM), substrate concentrations [443]. Subsequently it has been shown that lamB mutants have a 100 to 500-fold increased K m for maltose transport and that they cannot take up maltotriose [448]. Thus, Szmelcman and Schwartz concluded that the X-receptor facilitates the diffusion of maltose and maltotriose through the outer membrane [448]. It has been thought that the X-receptor pore was specific for maltose and maltodextrins. The first indication that the X-receptor also facilitates the diffusion of other nutrients came from Von Meyenburg and Nikaido [449]. More recently, in vitro experiments have shown that it behaves as a general pore [351,428,450,451]. The observed binding of maltose and of certain related compounds to the X-receptor pore [351,452--455] probably explains the relatively high permeation rate of these solutes through this pore. As maltodextrins too large to be transported are also bound by the outer membrane [453] the binding site must be located at the cell surface. Obviously phage lambda does not interfere with the binding site as maltose does not inhibit the rate of adsorption (M. Schwartz, personal communication). Interestingly, evidence for a cooperation of the lambda receptor with the periplasmic maltose binding protein in the uptake of maltose, as first hypothesized by Endermann et al. [456], has recently been obtained [442,457,458]. This interaction may also contribute to the preference of the pore for certain solutes. It has been shown recently that among lamB missense mutants strains occur with different in vitro pore properties and with poor in vitro interaction with maltose binding protein [449]. Such an approach will contribute to our understanding of the molecular functioning of the LamB protein,

The lambda receptor protein has been purified

TA

BL

E I

ll

E.

CO

LI

K-1

2 P

OR

E P

RO

TE

INS

NO

T A

NT

IGE

NIC

AL

LY

RE

LA

TE

D T

O P

EP

T1

DO

GL

YC

AN

-AS

SO

CIA

TE

D G

EN

ER

AL

DIF

FU

SIO

N P

OR

E P

RO

TE

INS

"

Pro

tein

C

on

dit

ion

s fo

r op

- M

r (P

art

of)

rece

ptor

for

S

truc

tura

l ge

ne

Pro

po

sed

fun

ctio

n F

urt

her

cha

ract

eris

tics

ti

mal

exp

ress

ion

ph

age/

coli

cin

Pha

ge T

6 C

o-re

gula

ted

wit

h nu

- 26

000

T6,

Co

l K

ts

x, r

ain

9.2

Up

tak

e S

ynth

esis

cat

abol

ite

repr

essi

- re

cept

or

cleo

tide

tra

nsp

ort

nu

cleo

side

s an

d

ble

[436

];

incr

ease

d am

ou

nts

syn

- [4

22]

deox

ynuc

leos

ides

th

esiz

ed i

n (y

tR a

nd

deo

R m

u-

[422

,423

] m

uta

nts

[42

2];

up

to

abo

ut

4.1

04

co-

Pha

ge 2

, P

rese

nce

of

mal

tose

47

392

[424

] )~

[42

5];

KI0

[42

6];

lam

B,

rain

91.

0 U

pta

ke

rece

ptor

T

PI

[383

]; T

P5

[384

];

mal

tod

extr

ins

SS

l [3

891

But

B p

rote

in

Vit

amin

BI2

60

000

[430

] B

F23

, E

-col

icin

s,

btuB

, ra

in 8

9.0

Up

tak

e li

mit

atio

n [4

29]

Co

la

[384

] a

vita

min

BI2

C

ir p

rote

in

Fe

3 + l

imit

atio

n [4

32

}

7400

0 C

oil,

Col

V

cir,

min

44

Up

tak

e co

mp

lex

ed F

e 3+

?

Fh

uA

pro

tein

F

e 3÷

lim

itat

ion

[43

2]

7800

0 T

1, T

5, 1

~80,

Co

lM

fhuA

, m

in 3

.4

Up

tak

e F

e 3+

ferr

ich

rom

e F

ec p

rote

in

Pre

senc

e o

f ci

trat

e 80

500

--

fec,

min

7

Up

tak

e F

e 3÷

- [4

33,4

34]

citr

ate

Fep

A p

rote

in

Fe

3÷ l

imit

atio

n [4

32

] 81

000

Co

lB,

Co

lD

fep,

rai

n 13

U

pta

ke

Fe

3+-

ente

roch

elin

83

K p

rote

in

Fe

3+ l

imit

atio

n 83

000

--

Un

kn

ow

n

Up

tak

e

com

ple

xed

Fe

3+ ?

pies

can

be

pre

sen

t p

er c

ell.

Syn

thes

is i

s ca

tabo

lite

rep

ress

i-

ble

[427

];

up

to

105

copi

es p

er

cell

can

be

pre

sen

t; p

ore

dia

met

er 1

.5

nm

[42

8]

Syn

thes

ized

in

redu

ced

amo

un

ts i

n pe

ril

mu

tan

ts [

238,

239]

an

d i

n he

p-

tose

-les

s L

PS

mu

tan

ts [

237]

U

p t

o 2

00

-30

0 c

opie

s pe

r ce

ll [4

311

Syn

thes

is o

f C

ir,

Fep

A a

nd

83K

pro

tein

s is

red

uced

in

perA

mu

- ta

nts

[24

0].

" Se

e su

bsec

tion

III

E-6

.

8()

to homogeneity [319,351,408,441,450,456,460]. As first found in Schnaitman's laboratory (Schnait- man, C.A., personal communication) it can be isolated associated with peptidoglycan. Applica- tion of this procedure has been reported to result in large losses [428,456] but in our hands a slight modification of the method used for purification of general pore proteins yields essentially pure Lamb protein in a high yield [319]. The active form is a trimer [461]. The protein has neither chemical [456] nor immunological [319] similarity with other major outer membrane proteins. The observation that both the )~-receptor protein and OmpF protein are recognized by the same bacteriophage, TP1 [383], therefore is most likely explained by assuming that the phage uses different regions of its tail for recognition of the two proteins. In whole cells the protein is also accessible to antibodies [462].

From the recently established nucleotide sequence of the cloned lamB gene [424] a polypeptide of 421 amino acids with two cysteine residues can be predicted. Many amino acids are charged, mostly negatively, and long peptides without charged residues do not occur. Interest- ingly, the residues can be alligned such that many negative charges are neutralized by positive ones [424]. The first results of the promising study of the structure-function relationship of the protein have recently been published [463,464].

lllE-6c. Outer membrane protein involved in the uptake of vitamin BI2 (Table II1). Vitamin B12, M r 1327, is too large to pass the outer membrane through general pores and it therefore requires a specific outer membrane protein for facilitating its translocation across the outer membrane (for a recent review see Ref. 465). The btuB product, M r 60 000, has been purified and might be a glycopro- tein [430] as it was shown to be sensitive to peri- odate treatment. However, as such a treatment can also result in protein modification [466] the data should be interpreted with care. The protein is the receptor for phage BF23 and the E-colicins. The vitamin binds to the protein to the extent that it protects the cell from killing by colicin and phage [28,467], Binding in vitro can be obtained in the presence of LPS (C. Bradbeer, cited in Ref. 465).

lllE-6d. Outer membrane proteins involved in the uptake of ferric ions (Table III). The solubility

of ferric ions is extremely low (approx. 10 ~ M) and bacteria have developed systems to take up these essential ions by first complexing the ferric ions with low molecular weight chelators known as siderophores of siderochromes. (For excellent recent reviews see Refs. 468-470), Chelators used by Enterobacteriaceae are aerobactin, enterochelin (enterobactin), citrate and ferrichrome. Interest- ingly, the antibiotic albomycin is a structural ana- log of ferrichrome [471,472]. In addition to these high affinity uptake systems, ferric iron can be taken up by a low affinity, chelator-independent system [465]. Alternatively, the uptake can be explained by low constitutive levels of the high affinity system which are present irrespective of the cultural conditions [468].

E. coli grown under conditions of limiting ferric iron derepresses the synthesis of siderophores and of several outer membrane proteins from which several have been identified as proteins involved in the uptake of specific iron-chelator complexes (see Table III). (For a recent review see Ref. 470). In at least one case it is likely that substrate recognition plays a role in uptake as it has been shown that the rate of adsorption of bacteriophage T5 is inhibited by the presence of ferrichrome [473]. The production and uptake of siderophores can play important ecological roles, e.g. it can be an important virulence factor [474-480] whereas siderophore production can be beneficial for plant growth by inhibiting growth of plant pathogens [481]. Siderophore production is temperature sensitive [482] and it therefore has been suggested that fever might be a host defence mechanism designed to deprive the pathogen of iron [482].

IIIE-7. Characteristics of E. coli outer membrane proteins without identified function

lllE-7a. Protein a. In E. coli K-12 protein a (M r approx. 40000, Ref. 205) usually is present in low amounts compared with other major outer membrane proteins [95,205]. It is hardly detected after growth at 30°C but rather high amounts can be produced at 42°C [95,484]. As protein a was not detected among the membrane proteins in the M r range of 30000 to 42000 in any of 45 E. coli hospital isolates [301], the production of large amounts might be typical for strain K-12. Strong support for this idea comes from recent experi-

ments which show that interactions between an 81 kDa ferric enterobactin pore protein and protein a (see further on) is confined to strain K-12 of E. coli [485].

Gayda and Markovitz have cloned the structural gene for protein M2, which they identified as protein a [486]. They claim that subsequent studies with plasmid mutants demonstrated that this protein is important in repressing the synthesis of capsular polysaccharide [487]. Earhardt et al. [488] have described deletion mutants which lack protein a. Their observation that such strains are normal with respect to capsule formation [488] is not consistent with the proposed role [487] of protein a in repressing capsular synthesis. The data of Earhardt et al. suggest that the structural gene for protein a is located at rain 12.5 of the chromosomal map [488]. These authors have also isolated a phage, LP81, which was claimed to require protein a for infection [488]. However, it has recently been shown that its real receptor is not protein a but a protein whose structural gene is located close to that of protein a (C.F. Earhardt, personal communication).

The outer membrane receptor for ferric enterobactin, FepA protein with a M r of 81000, is converted to a M r 74000 protein by an outer membrane protease, which has chemical and physical properties ascribed to protein a [248]. Moreover, as mutants lacking protein a do not convert the FepA protein [248] the evidence for protein a being a protease looks rather good. In fact it resembles the signal peptidase in this enzymatic activity, in its apparent molecular weight and its localization in the outer membrane (see subsection IIIE-2). It seems worthwhile determin- ing whether these two proteins are identical.

l l lE-7b. Protein I lL This protein (M r 17000, Refs. 222 and 489), which probably is identical with protein G [490,491] is a major outer membrane protein which occurs in l 0 4 t o 10 5 copies per cell [294]. The search for the function of protein III is hampered by the fact that no mutants have been found which lack the protein. Outer membranes of E. coli K-12 deep rough LPS mutants contain decreased amounts of protein III [93]. A protein band in the electrophoretic position of protein III was observed in all Enterob- acteriaceae mentioned in Ref. 322 (Lugtenberg, B., unpublished data).

81

As the amino acid compositions of protein III [294,295] and type 1 pilin [53] are very different, the possibility that protein III could be identical to pilin, as suggested by McMichael and Ou [492], can be ruled out.

Wu and Heath [493] reported that all LPS of E. coli is covalently linked to a protein with a mini- mum M r of about 14000, for which protein III was an obvious candidate. Indeed, large amounts of LPS are found in purified preparations of protein III but this co-purification is coincidental as it has been described that both components can be purified free from the other by using mild procedures [295]. Similar results were found in other laboratories and it is now generally agreed upon that LPS is not covalently bound to any protein.

With respect to the regulation of the synthesis of protein III it is interesting to note that it has been reported that its synthesis is negatively controlled by cyclic AMP at the level of transcription by a mechanism which involves the cAMP receptor protein [494]. An alternative interpretation is that cAMP controls the synthesis of OmpF protein and that changes in the level of protein III (and of OmpC protein) are simply responses to the level of OmpF protein [405].

IIIE-7c. LPS binding protein. The purification of a LPS binding protein from Re mutants of S. minnesota has been described. This M r 15 000 protein is surface located and can be considered as a common antigen in Enterobacteriaceae [495]. Al- though several properties suggest a possible identity with protein III, such a relation is not very likely as Re mutants produce little protein III [93] and as the amino acid compositions of the two proteins, especially the relative amounts of proline, glycine and tyrosine, differ too much [295,495]. Similarly, the amino acid composition excludes an identity with lipoprotein (compare e.g. Refs. 222 and 495).

IllE-Td. Outer membrane proteins induced by sulphate limitation. Growth of E. coli K-12 at Na2SO 4 concentrations below 50 ~M results in induction of the synthesis of two cell envelope proteins of M r 15 000 and 19 000. Preliminary experiments indicate that the M r 19000 protein and probably also the M r 15 000 proteins are located in the outer membrane (Lugtenberg, B. and Over- duin, P., unpublished data).

82

lllE-Te. Phage- and plasmid-coded outer membrane proteins. Phage lambda codes for an outer membrane protein (M r 20 500) for which no function has been established so far [328]. Upon infection of E. coil with the virulent phages T4 and T2 several new proteins of unknown function appear in the outer membrane thereby inhibiting the production of the major outer membrane proteins of the host [496,497].

The F-factor, a plasmid present in the best known E. coli donor strains in conjugation, codes for a series of outer membrane proteins [8]. Of these the, TraT protein has been studied most extensively. This protein ( M r 25 000) is involved in surface exclusion i.e. it prevents a donor cell from functioning as an acceptor cell by blocking the stabilization of mating aggregates [6]. It has been proposed that several specific receptor molecules are involved in the initial stages of conjugation and that the action of the TraT protein in surface exclusion is due to specific binding to one of these donor or recipient receptor molecules in the outer membrane [332]. In a detailed study Manning et al. [498] have shown that TraT protein is peptidoglycan-associated and exists as multimers in the outer membrane. No pore activity was detected. lts isoelectric point is very basic (pH approx. 9). Embedded in the membrane, the protein is resistant to trypsin. In whole cells it is exposed to the cell surface as it is accessible to CNBr-activated dextran and it can strongly be labeled with 125I, using lactoperoxidase [498]. Interestingly, the TraT protein is responsible for the resistance to the bactericidal activity of normal serum towards E. coli carrying plasmid R6-5 [499,500], R100 [501] or R222 [502]. In the latter case the authors suggest that the protein, named MRB protein, has an as yet unidentified transport function [502].

IV. Molecular organization of the outer membrane

I VA. Introduction

Microbiologists have realized already long ago that the envelope of Enterobaeteriaceae has very peculiar permeability properties. On one hand these bacteria are among the fastest growing cells in nature and they therefore must be able to provide the biosynthetic machinery inside the cell

with nutrients from the medium in an extremely efficient way. On the other hand, as growth of these bacteria is not affected by the presence of high concentrations of bile salts, detergents and many antibiotics, these cells must have an efficient barrier for the latter agents and it has been realized early that the outer membrane constitutes this barrier. As detergents and many antibiotics are hydrophobic whereas nutrients are hydrophilic, Nikaido has proposed two possible pathways for solute permeation through the outer membrane, a hydrophobic one and a hydrophilic one. In order to explain the discriminatory role of the outer membrane with respect to the permeation of solutes, he proposed that the hydrophobic pathway is non-existent in Enterobacteriaceae but does occur in some other Gram-negative bacteria in the form of phospholipid bilayer regions [361]. The hydrophilic pathways were assumed to consist of aqueous or water-filled pores in the membrane through which nutrients permeate, largely by a diffusion-like process (see subsection IIIE-5).

The permeability properties of E. coli cells can be altered by treatment with EDTA under conditions which do not influence the viability of the cells. This results in the release of half of the cellular LPS, in a strong increase in the permeability for detergents and hydrophobic antibiotics and in accessibility of cellular phospholipids for exogenous phospholipases [ 127,161 ]. The damage can be restored upon subsequent incubation in fresh medium [85].

The permeability of the cell can also be increased by mutations which lead to a deficiency for one or more outer membrane proteins or which reduce the length of the LPS sugar chain. Espe- cially the so-called deep-rough mutants of the Re and Rd chemotypes (Fig. 3) have a largely increased sensitivity towards hydrophobic compounds [410,463,503-505]. LPS mutants as well as an overwhelming number of mutants deficient in one of the outer membrane proteins appeared to be very useful in topological studies. It should be noted, however, that the lack of a constituent brings about a reorganization in the molecular make-up of the membrane. Several examples of such a reorganization are known. The lack of major proteins is compensated for by increased amounts of other proteins [96,232,506], of LPS

TA

BL

E

IV

TO

POG

RA

PHY

O

F O

UT

ER

M

EM

BR

AN

E

PRO

TE

INS

Prot

ein

Pres

ence

at

th

e Pr

otei

n-pr

otei

n ce

ll su

rfac

e in

tera

ctio

ns

Prot

ein-

pept

idog

lyca

n in

tera

ctio

ns

Prot

ein-

LPS

in

tera

ctio

ns

Lip

opro

tein

(fre

e fo

rm)

No

evid

ence

. Is

not

kn

own

as

a

rece

ptor

fo

r a

phag

e.

Doe

s no

t in

tera

ct

with

ex

ogen

ous

CN

Br-

dext

ran

or

with

an

ti-lip

o-

prot

ein

seru

m

(512

1.

Lip

opro

tein

(bou

nd

form

) N

o ev

iden

ce.

Om

pA

prot

ein

Rec

epto

r fo

r ph

ages

[2

93.3

81];

re

actio

n w

ith

antis

erum

[5

17];

the

anal

ogou

s 33

kD

a pr

otei

n of

S.

typh

imur

ium

re

acts

w

ith

bact

erio

cin

4-59

.

Om

pC,

Om

pD,

Rec

epto

r fo

r ph

ages

(s

ee

Tab

le

Om

pF

and

PhoE

II

);

reac

tion

with

an

tibod

ies

prot

eins

13

41 a

nd

CN

Br-

dext

ran

[519

].

Lam

B

prot

ein

Rec

epto

r fo

r ph

ages

(s

ee

Tab

le

III)

; re

actio

n w

ith

antib

odie

s

[462

].

Hom

olog

ous

olig

omer

s,

mai

nly

dim

ers,

ar

e fo

rmed

up

on

chem

ical

cros

s-lin

king

[S

131

. Can

al

so

be

cros

s-lin

ked

with

pr

otei

n II

I

[514

], an

d w

ith

Om

pA

prot

ein

to

a di

mer

ic

com

plex

[5

13].

Evi

denc

e fo

r in

tera

ctio

n w

ith

gene

ral

pore

pr

otei

ns

in

vitr

o

1515

1.

Evi

denc

e fo

r in

tera

ctio

n w

ith

gene

ral

pore

pr

otei

ns

in

vitr

o

[516

].

Can

be

ch

emic

ally

cr

oss-

linke

d to

hom

olog

ous

olig

omer

s [5

13.

5141

and

to

a he

tero

logo

us

dim

er

with

the

free

fo

rm

of

lipop

rote

in

[513

].

Can

be

ch

emic

ally

cr

oss-

linke

d to

di-.

tri-

, he

xa-

and

nona

mer

s

[398

,399

.513

,518

,520

] bu

t no

t

to

lipop

rote

in

[397

,399

,513

.520

,

5211

; bi

olog

ical

ly

activ

e fo

rm

is

trim

er

[356

,357

].

Can

be

ch

emic

ally

cr

oss-

linke

d to

di-,

tri-

, he

xa-

and

nona

mer

s

1461

1; t

rim

er

is b

iolo

gica

lly

activ

e fo

rm

1524

1.

Can

be

ch

emic

ally

cr

oss-

linke

d to

pept

idog

lyca

n [5

131

Cov

alen

tly

boun

d to

pe

ptid

o-

glyc

an

[265

.266

];

infl

uenc

es

pore

prot

ein-

pept

idog

lyca

n in

ter-

actio

ns

in v

itro

[5 1

61;

such

an

inte

ract

iion

is u

nlik

ely

in v

iva

1272

1.

Can

be

ch

emic

ally

cr

oss-

linke

d to

pept

idog

lyca

n 12

34.5

14.

5 18

1.

Can

not

be

cros

s-lin

ked

to

pep-

tidog

lyca

n [5

13,5

18];

in

vitr

o

reco

nstit

utio

n of

co

mpl

exes

of

Om

pF

prot

ein

[522

,523

] an

d

Om

pC

prot

ein

[516

] w

ith

pept

i-

dogl

ycan

-lip

opro

tein

.

In

vitr

o re

cons

titut

ion

of

com

-

plex

es

of

Lam

B

prot

ein

and

pep-

tidog

lyca

n-lip

opro

tein

[5

25].

No

evid

ence

.

No

evid

ence

Prot

ein-

LPS

co

mpl

ex

is r

ecep

tor

for

phag

es

K3

[293

] an

d T

uII’

[381

] an

d fo

r do

nor

cells

in

F-fa

ctor

m

edia

ted

conj

ugat

ion

[293

.308

];

com

plex

es

also

oc

cur

in

vivo

as

evi

denc

ed

by

phag

e

adso

rptio

n ch

arac

teri

stic

s [1

70]

and

free

ze-f

ract

ure

stud

ies

[200

].

Free

ze

frac

ture

st

udie

s in

dica

te

prot

ein-

LPS

co

mpl

exes

[Z

OO

].

Hep

tose

-les

s m

utan

ts

have

stro

ngly

de

crea

sed

amou

nts

of

Om

pF

prot

ein

190.

94)

and

PhoE

prot

ein

[236

] in

th

eir

oute

r m

em-

bran

e.

Pore

ac

tivity

(3

261

and

phag

e re

cept

or

activ

ity

[217

,226

,

3811

in

vitr

o ar

e de

pend

ent

on

LPS

.

Dec

reas

ed

amou

nts

of

lam

bda

rece

ptor

ar

e pr

esen

t in

he

ptos

e-

less

L

PS

mut

ants

[2

37];

ce

rtai

n C

%

part

icle

s ar

e pr

obab

ly

Lam

B

prot

ein-

LPS

co

mpl

exes

[2

00].

84

[93,127,129] and of phospholipids [88,127,129]. In the latter case the mutation most likely results in the insertion of part of these phospholipids into the outer monolayer, thereby making the cells sensitive towards detergents and exogenous phospholipases [96,127]. Defects in the structure of LPS, especially those caused by deep-rough mutations, can result in decreased amounts of outer membrane proteins [87,90,92,96] and in increased amounts of phospholipids [87,88,92,93] and LPS [92]. The resulting outer membranes are schematically represented in Fig. 6. A more subtle example of the mentioned reorganization is that substantial amounts of OmpA protein, which cannot be isolated complexed with peptidoglycan from E. coli K-12 wild-type cells, are found associated with peptidoglycan in pore protein deficient mutant cells (Lugtenberg, B. and Van Boxtel, R., unpublished data). Finally, mutant cells with heptose-less LPS or lacking Braun's lipoprotein excrete outer membrane blebs, which (at least in wild-type cells) have an architecture different from their cell-bound membrane in that the relative amounts of several proteins are drastically altered [272]. Thus, it is evident that mutants should be used with care for topological studies.

The cell surface of E. coli was investigated using freeze etch electron microscopy [507] and found to contain numerous randomly spaced depressions of about 4.5 nm in diameter, which could be the entrances of the aqueous pores [200,507]. Freeze- fracture studies were applied to study the interior of the outer membrane. It was shown that the outer membrane can be cleaved into two halves [88,508-510] suggesting a lipid bilayer structure. The two leaflets clearly differ in that the concave or outer fracture face (O~M) is covered with particles, 4 to 8 nm in diameter, whereas the O~M, the convex or inner fracture face, contains pits, probably complementary to the particles. Particles and pits are considered to be reflections of protein-LPS interactions (Refs. 2, 15,200, 511; see also subsection IVG-2). As it has been proposed that a substantial part of the particles represents aqueous pores [363], it is interesting to note that EDTA treatment reduces both the number of derepres- sions at the cell surface [507] as well as the number of intramembranous particles and pits [200].

In the following section we will describe the

available data on the molecular organization of the outer membrane, as mainly studied in E. coli and S. tvphimurium. Studies on the surface localization of outer membrane constituents indicate that LPS and almost all proteins, but not the phospholipids, have sites exposed at the surface which can act as receptors for phages, colicins and donor cells in conjugation and can react with antibodies, enzymes and chemicals. In addition, protein-protein and protein-peptidoglycan interactions of the major proteins are described. The organization of the phospholipids and LPS in the lipid bilayer and the way the planar structure of this bilayer is disturbed by protein-LPS complexes, visible in freeze-fracture electron microscopy as particles with complementary pits, will be treated in subsection IVG-2. Finally, a molecular model is presented which contains the knowledge collected up to now.

1 VB. Methods used for studying the localization of individual outer membrane constituents and their interactions

I VB-I. Localization at the cell surface One can conclude that a certain protein has a

site exposed at the cell surface if it can be shown that the protein is the receptor for a phage or a bacteriocin. This seemingly simple approach is hampered by the fact that in vitro experiments usually show that the receptor activity is lost upon purification of the protein, often because of a complex of both protein and LPS is required for receptor activity (see Table IV and subsection IVG-4). Therefore the protein is not necessarily the first recognized, and therefore surface-exposed, part of the receptor. Only in the case of OmpA protein strong evidence has been presented for a primary role of the protein by showing that the deeply burried lipid A portion is the active component of the LPS (Refs. 171 and 526). In all other cases one should be careful not to over-interprete the receptor evidence. However, the observation that resistant mutants often lack the protein and usually do not bind the phage or bacteriocin any- more shows that the protein is the most likely candidate for the primary receptor and that phages and bacteriocins can be used as rather reliable indicators of surface localization. According to

this criterium, many outer membrane proteins are recognized by phages and colicins (see Tables II-IV, and subsection IVG-4).

Proteolytic enzymes have been used as tools for demonstrating surface localization of proteins. In intact cells of E. coli K-12 none of the outer membrane proteins were degraded by proteolytic enzymes [527], which is not so surprizing as the natural habitat of Enterobacteriaceae is rich in such enzymes.

Membrane-impermeable CNBr-activated dextran has been used as a tool to couple surface-exposed outer membrane constituents [519]. In the case of S. typhimurium this approach showed that the peptidoglycan-associated general pore proteins are surface-exposed. Unfortunately, it is not possible for us to interprete the experiment carried out with E. coli strain B [519] with certainty. For example, the authors describe that a M r 31000 protein is exposed which they presume to correspond with protein III [519] although the latter protein has a M r of about 17000.

Several radio-iodination techniques have been applied to whole cells. In the case of the lactoperoxidase method it has been reported that most outer membrane proteins of E. coli K-12, including both OmpA protein and protein I, can be labeled [528]. We have obtained similar results but as we calculated that only a few protein molecules per cell were labeled, the labeling can also be explained by outer membrane damage of a small fraction of the cells (Van Alphen, L. and Lugten- berg, B., unpublished data). Munford and Gotsch- lich [529] have clearly shown how careful one must be with the interpretation of radio-labeling techniques. Using the chloramine T method they found that under certain conditions only the lipoprotein was labeled which contains only one tyrosine residue located only three amino acid residues from the covalent linkage of the lipoprotein with the peptidoglycan [529]. Obviously protein pores are permeable for chloramine T. Recently promising results with non-Enterobacteriaceae have been obtained with the radio-iodination reagents IodoGen (1,3,4,6-tetrachloro-3 a,6 a-diphenyl glycoluril) [530,531] and DISA (diazotized [35S]sulfanilic acid) [103,532].

An obvious method to show surface localization is the detection with mono-specific antibodies at

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the cell surface, as has been used for ECA (see subsection IVE) and LPS (see subsection IVC), in the cases of E. coli LamB protein [430] and OmpA protein [517] and peptidoglycan-associated general pore proteins of S. typhimurium [34].

IVB-2. Protein-protein and protein-peptidoglycan nearest neighbour associations

A variety of bifunctional cleavable cross-linkers with various specificity and distances between the reactive groups can be used to show interactions between membrane constituents [533]. This approach has been applied successfully in the case of outer membrane proteins (see Table IV). It should be noted that when two molecular species cannot be chemically cross-linked the result not necessarily means that these constituents are not neighbours as the spacing might not have been correctly chosen or as reactive groups might not be present in the right positions (e.g. see subsection IVF-4). Moreover, it should be realized that proteins which meet each other only for a short time during the experiment can also be cross-linked. The latter type of cross-linking due to random collisions can be minimized by using photo-sensitive reagents which can be activated for a short time (approx. 1 ms) with a flash [533]. In general, one should be careful in interpreting crosslinking data. Selective cross-linking with affinity labels is preferable to nonselective cross-linking (for instance with bis-imido-esters). The latter reagents are useful for the detection of interactions, provided that various reagents are applied under well-defined conditions, such as short reaction times, low concentrations of reagents to avoid extensive cross-linking due to random collisions and defined temperatures to account for phase separation of membrane components. The physiological significance of the complexes generated by cross-linking can only be established by demonstrating that the complexes consisting of the same membrane components as observed after cross-linking are required for biological activity .In this respect the homotrimers of pore proteins seem to fullfil these requirements (see subsection IVF-4).

Some outer membrane pore proteins can be isolated tightly complexed to the peptidoglycan (see Tables II and III). The question of whether such complexes also exist in vivo will be discussed in subsection IVF-5.

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1 VB-3. Interactions between individual proteins and LPS (see also subsection 1 VG-4)

LPS is often required for in vitro biological activities of outer membrane proteins, e.g. receptor activity for phages, for donor cells in conjugation, for a tight association with peptidoglycan and for pore activity. Strong evidence that OmpA protein- LPS interactions also occur in vivo has been provided [170]. Some proteins are hardly or not at all found in the outer membrane of heptose-less LPS mutants. The application of freeze-fracture and biochemical techniques on appropriate mutants has indicated the existence of protein-LPS interactions for many major proteins (see subsection IVG-2). Finally, recent experiments have shown that passive immunization of animals with antibodies raised against protein-LPS complexes, from wich the anti-LPS activity had subsequently been removed, results in a much better protection against experimental infection than the use of antibodies raised against highly purified protein (Refs. 534 and 535; Dankert, J., Hofstra, H. and Veninga, T.S. (1980) in FEMS Symposium on Microbial Envelopes, Saimaanranta, Finland, Ab- stract 51).

1 VB-4. The lipid matrix and interactions of proteins

with lipids X-ray diffraction, 31p-nuclear magnetic reso-

nance spectroscopy (31P-NMR) and freeze-fracture electron microscopy are standard methods for studying the organization of membranes, especially of its phospholipids, since extensive data are available from model studies. With X-ray diffraction the presence of regularly arranged fatty acyl chains in regions of a fair size can be detected, such that the occurrence of phospholipid and LPS in bilayers can be detected. Using 31p-NMR spectroscopy, which measures the ways of motion of phosphorus-containing components (including LPS), lamellar, isotropic and hexagonal phases were detected. This technique is especially powerful in combination with freeze-fracture electron microscopy, which reveals the morphology of the fracture faces. These faces are usually the sites where the ends of the fatty acyl chains approach each other. A special problem for the 31P-NMR study of outer membranes is the presence of LPS, which contains several phosphorus atoms, thereby

unequivalently attributing to the :~P-NMR spectra.

2H-NMR and electron spin resonance (ESR) spectroscopy are both useful for the study of interactions of the specifically labeled compounds with their environment. Since the time scale of the measurement is relatively long with NMR ( l0 4 l0 t cm 2. s - l ) in comparison with ESR (10 v- 10 ~ cm ~- s ~), long-term and short-term interactions can be discriminated.

1 VC. Localization of LPS

Studies in which ferritin-labeled antibodies directed against the O-antigenic part of the lipopolysaccharide of S. typhimurium were used, have clearly shown that LPS is located exclusively at the outside of outer membranes [36]. This conclusion was confirmed recently, since it appeared that the sugar chains of the LPS molecules in whole cells of S. typhimurium are oxidized by exogenous galac- rose oxidase [536]. LPS occurs in various forms in the outer membrane. Half of it can be extracted with EDTA [199]. Newly synthesized LPS does not mix with old LPS [537], indicating different domains. Finally, outer membrane proteins form strong complexes with a substantial portion of the LPS molecules (see subsection IVG-4).

1 VD. Localization ofphospholipids

Phospholipids are part of the lipid bilayer of the outer membrane (see subsection IVG). As it has been calculated that the outer membrane contains hardly enough phospholipids to cover one monolayer [88] and as it has been established that LPS is located exclusively at the outside of the outer membrane (see subsection IVC), it seems likely that phospholipids are mainly or completely located in the inner monolayer. The fact that, despite serious attempts, investigators have not succeeded in showing accessibility of phospholipids by exogenous agents in intact cells of E. coli strains K-12 and B, and of S. typhimurium, sup- ports this idea. Perhaps even more convincing is the simple observation that wild-type cells and Rc mutants can grow in the presence of bile salts and SDS, whereas Re mutants and pore protein deficient mutants are sensitive to these and other

hydrophobic agents [503-505,538]. Cells of the sensitive strains have increased amounts of phospholipid [88,92,93,127], which are probably located in the outer leaflet (Refs. 88 and 127; subsection IVA). Such areas of phospholipid bilayer are likely to be susceptible to detergents. Similarly, chemical labeling of S. typhimurium cells with membrane- impermeable CNBr-activated dextran revealed that phospholipids are not accessible in wild-type cells and Rc LPS mutants but that phosphatidylethanolamine in cells of Rd and Re strains, which lack most of the core sugars of LPS [539] and therefore contain decreased amounts of protein (see subsection IVA), could be coupled to this agent. Using conjugated dansylchloride, Schindler and Tauber [540] showed that phospholipids in Rc mutant cells of S. typhimurium are not accessible for this agent unless the cells had been pretreated with chelating agents like EDTA and Tris. The use of various exogenous phospholipases shows that the phospholipids of intact cells of E. col] strains K-12 [127] and B [541] and of S. typhimurium Rc mutants [539] are resistant to degradation by phospholipases A 2 and C, irrespective of the source of the enzyme, thereby excluding that lack of degradation is due to influences of lateral surface pressure [127]. Tris-EDTA treatment sensitized cells of E. col] B and S. typhimurium, but not those of E. col] K-12 [127,539,541]. Degradation of phospholipids was also observed in cells of deep-rough LPS mutants [ 127,539] and of mutants deficient in several outer membrane proteins [ 127]. These experiments convincingly show that phospholipids are not accessible in intact wild-type cells. They strongly suggest a localization of the phospholipids in the inner monolayer, but do not strictly exclude a localization outside this layer in such a way that the phospholipid molecules are shielded against the used agents by endogenous molecules. Indeed, it has been shown that phospholipids can be protected by LPS against solubilization by detergents [163,324]. Moreover, phosphatidylethanolamine and LPS have been shown to form close complexes [139]. However, electron spin resonance studies have clearly shown, both for outer membranes [542] and for model bilayers [542,543], that phospholipids and LPS are segre- gated into separate domains. These data clearly indicate that the vast majority, if not all, of the

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phospholipid molecules of the outer membrane are localized in its inner monolayer. Consistent with this conclusion is the result of our own work [ 170], which shows that wild type cells of E. col] K- 12 the receptor for phage K3, consisting of a complex of LPS with OmpA protein [293], does not contain phosphol ip id molecules in its immedia te neighbourhood, but that in pore protein-deficient mutants the presence of phospholipid in these domains can clearly be shown [170].

The results of studies on the localization of LPS and phospholipids and the changes in chemical composition resulting from mutations and EDTA treatment enable us to draw models of outer membranes of the cells discussed in the previous sections. Schematic representations of outer membranes of wild-type cells, protein deficient mutants, heptose-less LPS mutants and of EDTA-treated cells are shown in Fig. 6.

IVE. Localization of ECA

The ease with which anti-ECA antibodies can be adsorbed by S- and R-form bacteria suggests a localization of ECA at the cell surface [544,545]. The recent chemical purification of ECA in Mayer's laboratory enabled the investigators to localize ECA reliably. Using either ferritin-labeled or fluorescent monospecific antibodies, Rinno et al. [202] demonstrated the cell surface location of ECA. Interestingly, more ECA was detected in immunogenic than in non-immunogenic strains [202].

Nothing is known about protein-ECA interactions except that the level of ECA in E. col] K-12 is not altered by the absence of one or more of the proteins OmpA-, OmpC- and OmpF-protein (Mayer, H., unpublished data).

I VF. Localization and topography of outer membrane proteins

1VF-1. Introduction Most proteins of the outer membrane are rich

in fl-structure [230,295,317]. They are not solubilized from the membrane by extraction with buffers with high ionic strength or with EDTA [97, 199, 222,294], but solubilization requires the presence of detergents or strong chaotropic agents. Ionic

88

PL ~,~,~,~ ~ ~ ~ ~,~ ~ ~,~ ~ ,L wild type

K-12

pore protein deficient mutant

~ ~I~t~ 1~ ~ll '~ mutant missing c 1~ ~ 1 1 ~ ~ 1 1 I ~ ~I~I~ m. A

d ,~ ~ ~ 1 1 I ~ I ~ 1 ~ 1 ~ 1 1 1 ~ t ~ h ept o s e_le s s ~ J ~ l ~ , ~ ~ l ~ ~l~q~,~ LPS mutant

EDTA treated wild type

Fig. 6. Schematic models of the bilayer organization of the outer membrane of cells of (a) wild-type E. coli K-12 and Rc mutants of S. typhimurium, (b) a pore protein deficient strain, (c) a strain lacking the three major proteins OmpA, OmpC and OmpF protein, (d) a deep rough LPS mutant and (e) wild type E. coli K-12 treated with EDTA. The LPS molecules (L) are present exclusively in the outer leaflet (OL). In wild type E. coil K-12 (a) all phospholipid (PL) is drawn in the inner leaflet (IL) whereas for reasons of simplicity only transmembrane proteins (P) are shown. Outer membranes of cells with complete LPS are supposed to have the same structure except that the sugar chains are longer. In pore protein deficient mutants (b) part of the missing protein is replaced by molecules of another protein whereas the remainder is compensated for by additional amounts of LPS and phospholipid [127]. The latter compensa- tion is assumed to lead to insertion of phospholipid in the outer monolayer. The resulting phospholipid bilayer regions explain the observed sensitivity of such cells to SDS [236] but are obviously too small to be attacked by exogenous phospholipases [127]. Sensitivity to phospholipase is observed when, in addition to the two pore proteins, also OmpA protein is lacking. The outer membrane of such a strain (c) is very poor in transmembrane proteins, the lack of which is compensated for by an increase in both LPS and phospholipid content. Deep rough Re mutants (d) have outer membranes with extremely short LPS sugar chains (see Fig. 3). Substantial amounts of outer membrane proteins are lacking [90,92-94] which is compensated for by increased amounts of LPS and phospholipid, resulting in sensitivity to detergents [503,504], hydrophobic antibiotics [538] and phospholipases [127]. EDTA-treated E. coli K-12 cells (e) have lost half of their cellular LPS [85] but no protein [200]. The resulting lesions in the outer leaflet of the outer membrane are presumed to be rapidly filled with phospholipid molecules derived from the inner leaflet and from the cytoplasmic membrane, The resulting cells are transiently sensitive to hydrophobic antibiotics [85] and phospholipases [127,541]. For reasons of simplicity ECA and divalent cations have not been indicated.

interactions seem to be impor tant for their anchor-

ing in the membrane since either an ionic deter-

gent or a combina t ion of a non- ionic detergent

(e.g. Tr i ton X-100, Sarkosyl) with EDTA is required to solubilize the proteins [97,294,317]. Stud-

ies on the localization of outer membrane proteins are merely restricted to three types of major proteins, namely lipoprotein, OmpA protein and

peptidoglycan-associated pore proteins. The present knowledge on their localization will be described in detail.

I VF-2. The major lipoprotein (see also Table 1 V) One third of the total amount of l ipoprotein is

covalently bound to peptidoglycan, whereas the remainder exists in the free form. Evidence for a cell surface localization of either form is lacking.

Bacteriophages using the l ipoproteins as (part of) their receptor or coupling of the l ipoprotein from

intact cells to exogenous CNBr-activated dextran

have never been described. Moreover, ant iserum directed against the l ipoprotein does not react with

intact wild-type cells [512]. From X-ray analysis of paracrystals of the lipo-

protein, this protein appears to consist of a super- helix or a coiled coil structure [280]. Since a high content of a-helix and lack of #-structure was deduced from infra-red and circular dichroism spectra, long stretched a-helices are the most likely

conformat ions of the l ipoprotein [278,546,547]. Trea tment of peptidoglycan-outer membrane

complexes with chemical cross-linkers results in dimerizat ion of the free form of the l ipoprotein.

Moreover, free l ipoprotein can be linked to peptidoglycan and also to OmpA protein in a dimeric complex [513]. Evidence for another interact ion of

l ipoprotein, namely with pore proteins, comes from work which shows that the in vitro interaction between pept idoglycan and pore protein is weakened in a mutan t lacking l ipoprotein and by trypsin t reatment which cleaves the bound form of the l ipoprotein from the peptidoglycan [515]. Simi- larly, experiments designed to study the interactions between peptidoglycan and pore proteins have shown that the presence of bound lipoprotein is required to obta in an ordered lattice structure [375,398,516]. Also spin labeling studies provide evidence for an interact ion between l ipoprotein and pore proteins [548]. Therefore it initially was

surprising that no cross-linking was found between lipoprotein and OmpC or OmpF proteins [397,513,520,521] or between lipoprotein and Salmonella pore proteins [399]. More recent data indicate that the vast majority of the amino groups are located within the pore [550,551] and therefore are not available for cross-linking. Thus, the present results certainly do not exclude a direct interaction between free lipoprotein and pore proteins. For a discussion on this subject the reader is referred to subsection IVF-5.

Three different models for the localization of the lipoprotein have been proposed. Inouye [280] and McLachlan [552] propose an uninterrupted a-helix with a width of 7.6 nm and Braun [266] proposes two anti-parallel a-helical segments inter- rupted by a bend near the middle of the molecule. Recently, Inouye and co-workers have proposed a new model consisting of transmembrane assemblies of three monomers of the pore protein, each of which is associated with a trimer of lipoprotein [7].

Inouye and McLachlan propose interactions between at least six, and two lipoprotein molecules, respectively. The proposed existence of lipoprotein dimers has been confirmed by cross-linking studies [513,514] supporting McLachlan's model but not rejecting the others. Inouye's first model proposes a pore function for the lipoprotein [280]. Evidence for such a function could neither be obtained in vitro [2,338] nor in vivo [553]. Whereas in Inouye's models [7,280] the protein part spans the thickness of the outer membrane, it is located outside the lipid bilayer in the periplasmic space according to the outer models [266,552]. The latter models have the advantage that they leave some space between the outer membrane bilayer and the peptidoglycan layer. Under the usual physiological conditions this periplasmic space must be present between these two layers as it should be expected that due to turgor pressure no substantial space is left between the cytoplasmic membrane and the peptidoglycan layer. Moreover, Inouye's models which propose large transmembrane assemblies [7,280] are not supported by freeze-fracture electron microscopy experiments of wild-type cells and lpp mutants as no differences could be detected in the morphology of either of the outer membrane fracture faces (Van Alphen, L., Verkleij, A. and

89

Lugtenberg, B., unpublished data). The models of Braun [266] and McLachlan [552] propose that the acyl chains are anchored in the inner leaflet, whereas the protein traverses the periplasmic space. The observation that the protein is still integrated in the membrane of mutant cells lacking some or all of the acyl chains [554-557] seemingly con- tradicts these models. However, these results can be explained by an association of the protein part of the lipoprotein with OmpA protein or with pore proteins. In conclusion, the localization of the lipoprotein is far from certain. On the basis of the available experimental evidence we tend to favour the model of McLachlan. The outside surface of McLachlan's dimeric lipoprotein structure is hydrophilic, consistent with a location in the periplasmic space [552].

IVF-3. OmpA protein (see also Table IV) OmpA protein is an oligomeric transmembrane

protein. It has sites located at the cell surface (see Table IV). Its presence prevents cleavage of the two leaflets in freeze-fracture experiments [198,200,508]. It can be cross-linked to the free form of the lipoprotein and to peptidoglycan. The observation that ompA, lpp double mutants, but not the two single mutants, grow in spheres and lose part of their outer membrane by 'blebbing', strongly suggests that both the OmpA protein and the lipoprotein interact with peptidoglycan [271].

As intact subunits of OmpA protein (but not its pronase resistant fragments) could be cross-linked [513], it is likely that the interaction between individual OmpA protein molecules takes place near the C-termini in or near the periplasmic space. Evidence for a localization of the -COOH part of the OmpA protein molecule at the periplasmic side of the membrane comes from the observation that this part is only degraded by trypsin after damag- ing the cell such that the enzyme can reach the periplasmic side of the outer membrane (Refs. 213, 526, 527 and Van Alphen, L. and Lugtenberg, B., unpublished data). These experiments suggest a localization of the N-terminal part at the cell surface or in the membrane, whereas the C-terminal part could span the periplasmic space and even interact with the peptidoglycan. Consistent with this idea are the results on the localization of OmpA protein fragments lacking the C-terminal

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third of the molecule [310,311]. The transmembrane part is located between residues 1 and 177 and the hydrophobic segment of 27 residues is a good candidate to span the membrane ([269], see subsection IIIE-4).

I VF-4. Peptidoglycan-associated pore proteins (see also Table IV)

These proteins include those listed in Table II as well as the LamB protein. They have sites exposed at the cell surface as they interact with phages, antibodies and CNBr-dextran (Table IV). Experiments with chemical cross-linkers indicate that the proteins occur as trimers. Functional pore proteins of strains producing one pore protein species consist of three identical monomers [356, 357], which act in highly cooperative fashion (subsection IIIE-5). Despite attempts in several laboratories to convert inactive monomers into functional trimers renaturation has never been reported (see for instance Refs. 356 and 357).

Most of the advanced work on the structure of pore proteins has been carried out in Rosenbusch's laboratory using the matrix proteins of E. coli strain B E [230,373,375,558]. Electron microscopic examination of protein-peptidoglycan complexes of E. coli B ~ shows that the protein is arranged in a hexagonal lattice structure [230] with a repeat of 7.7 nm, which covers the outer surface of the peptidoglycan [558]. The periodic structure of such a two-dimensional crystal is maintained and even improved [375] in the absence of peptidoglycan, and is therefore based on strong protein-protein interactions [558]. A unit cell probably contains three protein molecules and a triplet of identations which may represent channels which span the protein monolayer [558]. Based on these and other observations a model of a pore trimer is proposed in Ref. 375. The thickness of the protein layer (4-5 nm) [375] is the same as that of the apolar part of a lipid bilayer (approx. 4.5 nm) suggesting that, if the structure still represents the in vivo situation, the porin is practically completely embedded in the membrane. Recently, three-dimensional crystals of matrix protein have been obtained [373,559,560]. Successful association into large crystals depended on the use of ~-octylg- lucoside [373]. The first results show that a large fraction of the polypeptide is present in fi-pleated

sheet structure with strands nearly parallel to the normal of the membrane plane [560]. It can be expected that this approach will yield considerably more structural information in the near future.

A beginning of a study of the structure-function relationship of OmpF protein was made in Nakae's laboratory. Isolated porins were chemically modified, incorporated into liposomes and tested for activity. The results suggest that hydrophilic amino acids play an important role in the diffusion of charged solutes through the pore [551]. More recently a similar approach has been undertaken in Rosenbusch's laboratory [550]. In the latter study electrophoretic mobility and several other parame- ters, but not pore activity, were used as criteria for native pores. The results support those of Nakae's laboratory [551] and seem to indicate that all but one of the lysine residues are contained within the pore [550]. Results with insertion and deletion mutations in the cloned genes ompF and phoE indicate that protein fragments are not easily obtained (see subsections IllE-5di and ii). The carboxy-terminal end of PhoE protein seems to be essential for the presence of the protein in the outer membrane [415], suggesting a situation different from that of OmpA protein from which the carboxy-terminus can be deleted without strong functional losses [310,311].

The question of whether heterologous or mixed pores, containing monomers of different pore proteins, exist in strains with more than one pore protein was approached in three different laboratories. Whereas the results of Palva and Randell [397] and of Ishii and Nakae [561] with E. coli K-12 and S. typhimurium, respectively, strongly suggest that mixed pores do not exist or only to a minor extent, results of Mizushima's laboratory indicate that when cells synthesize OmpC protein and OmpF protein simultaneously, heterotrimers are formed at random. In contrast, when the two proteins are synthesized in separate periods heterotrimers were not found [562].

The possible interaction of pore proteins with lipoprotein has already been discussed in subsection IVF-2.

Evidence for an interaction of pore proteins with LPS is summarized in Table IV and described in subsection IVG-4. It should be noted that only very little LPS can be cross-linked to pore proteins

91

of S. typhimurium [399], although it is known that pore proteins of E. coli interact with LPS [200,217,226,381].

Tight associations between pore proteins and peptidoglycan have been found in vitro. The question of whether such interactions also occur in vivo will be discussed in subsection IVF-5.

The LamB protein has many properties in common with the general pore proteins, but seems not to be related to these proteins [319]. It is surface- located (Table IV) and the native form of the lambda receptor is a trimer [461,563], which probably interacts with LPS [237,525], peptidoglycan [319,441,460,525] and with the bound form of the lipoprotein [525]. The lambda receptor activity is decreased in heptose-less LPS mutants [237], suggesting that the amount of this protein in the outer membrane of such mutants is decreased as has been described for other proteins [90,94-96,236]. Two-dimensional sheets recently obtained will allow structural studies [564].

Since anti-B-galactosidase reacts with intact cells containing hybrid proteins of the lambda receptor protein and fl-galactosidase [565], it has been proposed that the carboxy-terminus of LamB protein is located near the cell surface. As models for protein translocation are based on this result [5], it should be noted that the evidence for this assump- tions is insufficient [5]. The localization of the hydrophilic LacZ protein at the cell surface will be thermodynamically favourable but this not necessarily implies that the carboxy-terminus of LamB protein, without the extremely bulky LacZ protein, behaves in the same way.

IVF-5. Are matrix (pore)proteins associated with peptidoglycan in vivo?

Rosenbusch described that the insoluble fraction obtained after heating cells or cell envelopes of E. coli B E in 2% SDS at 60°C consists of peptidoglycan-lipoprotein complexes with non-covalently attached to it the OmpF protein organized in a regular so-called 'matrix' pattern [230]. This periodic structure is even improved upon removal of peptidoglycan [375]. Evidence has been presented that in vitro binding of the pore protein to the peptidoglycan occurs partly through the lipoprotein [515].

'Matrix' structures have been reconstituted in

Mizushima's laboratory. Initially it was shown that isolated OmpC protein and OmpF protein can be physically reassociated with peptidoglycan in the presence of 5 mM Mg 2+ [318]. Added LPS stimulates the binding, especially of OmpF protein [522]. Binding is not altered when the covalently bound lipoprotein has been removed from the peptidoglycan by pretreatment with pronase [318,522]. Electron microscopic examination showed that an ordered hexagonal lattice structure consisting of OmpC protein and LPS can be obtained over the entire surface of lipoprotein-bearing peptidoglycan [516], resembling that obtained after treatment of cells in 2% SDS at 60°C [230,558]. Omission of either protein or LPS resulted in failure of formation of lattice structure. No ordered lattice structure was formed when peptidoglycan without bound lipoprotein was used and OmpC protein and LPS assembled in vesicles [516]. Essentially the same results were obtained with OmpF protein. The role of LPS was studied in more detail and it was observed that when LPS was replaced by lipid A or even by fatty acid, an ordered lattice structure was also formed although the lattice constant was smaller [523,566]. The intriguing question of whether these structures are functional was studied using adsorption of phage T4 as a tool. Indeed, T4 phage adsorption onto reconstituted cell surfaces containing OrnpC protein as the protein component was observed. Both the protein and LPS were essential for this interaction. The needle even penetrated the cell surface [567] and, when phospholipid liposomes had been included in the system, the phage also ejected its DNA [567a]. In this case wild-type LPS could not be replaced by either heptose-less LPS or lipid A. A function is also attributed to the peptidoglycan in generating a flat surface large enough to interact with all tail fibres of a single phage particle [567].

The in vitro results described above raise the questions of whether the hexagonal (matrix) pattern is also present in vivo and even whether the observed association between pore proteins and peptidoglycan as well as the proposed association between pore proteins and lipoprotein mimicks the in vivo situation.

As argued by Rosenbusch et al. [375] it is unlikely that the regular array of pore proteins also exists in the native outer membrane but the

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hexagonal structure rather is a consequence of the extraction procedure [375]. The tendency of pore protein trimers to form regular aggregates on the peptidoglycan [230] and the improvement of the long-range order by peptidoglycan degradation [375] suggests the possibility that the protein-protein interaction between pore protein trimers might be an artefact which is easily obtained in a strain like E. coli B E with a number of monomers of as many as 105 per cell [230] or in reconstituted systems with highly purified constituents, but which is more difficult to obtain in strains containing several pore proteins or containing less pore protein. We have approached the question of the in vivo occurrence of regular patterns with the freeze-fracture technique which is so fast and simple that it is unlikely to give artefacts. Cleava, a,a~e planes through the outer membrane show OM particles which, as they are the reflections of protein-LPS aggregates (see subsection IVG-2), indicate the position of proteins. If the matrix pattern of the OmpF protein of the outer membrane E. coli strain B E described by Rosenbusch [230] also occurs in vivo, it should be observed by freeze fracturing of wild-type cells of E. coli B E, and even more striking in cells of an ornpA mutant of this strain. The choice of this mutant has the advantages that cleavage is preferential through the outer membrane in such mutant cells and that the OmpF protein represents over 90% of the outer membrane protein in this mutant. The results (Fig. 7A) show that, although the OM particles seem to be very homogeneous, no crystalline network is present. We consider this as the best evidence that the regular pattern is an in vitro artefact.

The answer to the question whether the association between pore protein with peptidoglycan, or in a trimeric complex which also contains the lipoprotein, is an artefact is much more difficult to give. The observation that the complex resists rather extreme conditions (2% SDS, 60°C) seems to be a strong indication for a natural interaction. Also the observation that the presence of LPS which is required for many biological activities of outer membrane proteins (see Tables II, III and IV and subsection IVG-4) increases the binding, suggests that it concerns a native situation. An indication of the opposite was the surprizing observation

that pore protein cannot [513] or hardly [518] be cross-linked to peptidoglycan. However, the observation that most charged amino acid residues seem to be located within the pore [550,551] is still consistent with a native complex as the pore protein probably has no reactive amino acid residues in the correct position for efficient cross-linking. Crystallographic data from Rosenbusch's laboratory [375] suggest that native pore protein only spans the apolar part of the membrane. This apolar part is probably at a distance of about 9 nm from the peptidoglycan due to the presence of the lipoprotein in the periplasmic space (see subsection IVF-2). It is hard to imagine that pore proteins are spanning the periplasmic space since no space would than be left for the periplasmic binding proteins which transport nutrients from the pore to the cytoplasmic membrane (see subsections IA and IIIE-5). The only experimental evidence for the absence of an association in vivo between pore proteins and peptidoglycan-bound lipoprotein complexes comes from analyses in Witholt's group of vesicles excreted in the medium by growing cells. In comparison with the corresponding cellular outer membranes these medium vesicles contain only 35% free lipoprotein and almost none of the bound lipoprotein. Medium vesicles also have reduced amounts of OmpA protein, which probably interacts with peptidoglycan, and of the bound form of the lipoprotein (Table IV), while they contain large amounts of pore forming protein 1 and LamB protein [272], suggesting that interactions between pore forming proteins and the peptidoglycan-lipoprotein complex - if existing are rather weak [272]. Summarizing, conclusive evidence for a direct interaction between peptidoglycan and pore proteins in vivo is lacking. An indirect interaction through lipoprotein molecules is conceivable but, if occurring, is certainly not operative for all molecules as discussed above [272]. Moreover, a space between peptidoglycan and pore proteins has the advantage that the functioning of periplasmic proteins is easier to imagine.

A schematic drawing of the topography of outer membrane constituents, as discussed so far in section IV, is presented in Fig. 8.

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E Fig. 7. Freeze fracture morphology of the outer membrane of an ompA mutant of E. coli strain B . The concave (outer) fracture face of the outer membrane (O"~I) is almost completely covered with particles (A) whereas the convex (inner) fracture face (OM) contains numerous pits (B), presumably complementary to the particles. In this case the particles are very homogeneous in size which presumably is caused by the fact that the vast majority of the particles and pits represent complexes of LPS with only one protein species, namely OmpF protein. Cells were slowly cooled from 37°C to 4°C in order to optimize conditions for the formation of twodimensional crystals. Subsequently the sample was quenched from 4°C. No signs of crystallization can be detected ( × 100,000). (From Verkleij, A., Leunissen-Bijvelt, J., Van Boxtel, R. and Lugtenberg, B., unpublished data).

IVG. The lipid matrix

1VG-1. Is the outer membrane a lipid bilayer? This quest ion was approached using three dif-

ferent techniques, namely X-ray analysis, freeze- f racture electron microscopy and 31p-NMR. Al- ready in 1975 it was repor ted that fat ty acyl chains of E. coli outer membranes occur in bi layer con- fo rmat ion as X-ray analysis of isolated membranes showed lamel lar reflections [128]. Freeze-f rac ture electron microscopy is widely used to s tudy the inside of membranes since fractur ing occurs between the fat ty acyl ends of the lipids, which is the

weakest par t of membranes [568,569]. Outer membranes of many Gram-nega t ive bacter ia are frac- t u r e d i n d e e d [ 8 8 , 2 0 0 , 3 0 8 , 5 0 8 , 5 1 0 , 5 1 1, 570-573], indicat ing that this membrane is basi- cal ly a bi layer [569]. Remarkab le is that only small fracture faces were ob ta ined in wi ld- type cells. This is obviously due to the nature of the outer membrane , since this phenomenon was also observed with isolated E. coli outer membranes [574]. The high content of t r ansmembrane proteins, especial ly the O m p A prote in [198,200], is likely to be responsible for this phenomenon. The outer f racture face of the outer membrane (OM) is

'~4

densely covered with particles, which consist of aggregates of protein-LPS complexes (see subsection IVG-2), thereby perturbing the planar lipid bilayer structure and making impressions (pits) in the inner fracture face (O~M) complementary to the particles (cf. subsection IVG-2). More recently, ~P-NMR was used to study the orientation of the phospholipid and LPS in the lipid matrix. The question was of special interest since at the growth temperature the isolated phospholipids yield 3L p_ NMR spectra and freeze-fracture morphology which are characteristic for a superposition of isotropic, hexagonal and lamellar phase [575], probably due to the presence of phosphatidylglycerol and diphosphatidylglycerol next to phosphatidylethanolamine [575]. The 31P-NMR spectra of the outer membrane (linked to peptidoglycan) turned out to be the superposition of lamellar phospholipids and lamellar IPS [162,574], identical to the spectra obtained from phospholipid-LPS liposomes, which only showed bilayer freeze-fracture morphology [162]. Moreover, using X-ray diffraction on purified LPS, lamellar reflections were observed, indicating that LPS itself is able to maintain the bilayer configuration [577]. In addition to the lamellar phase a small amount of non-bilayer phase was observed in ~L P-NMR spectra of outer membranes at and especially above the growth temperature [574], which may be related to translocation of proteins, LPS and phospholipids from the cytoplasmic membrane to the outer membrane (compare with subsection IC and Fig. 2). Although the amount of lipid participating in non-bilayer phases is small (less than 1%) in the time scale of the measurement, it is possible that a substantial amount of lipid is participating for a very short period of time, which would be consistent with the idea of temporary adhesion zones involved in the biogenesis of the outer membrane (Fig. 2). Four physico-chemical characteristics of the outer membrane might favour non-bilayer phases, since they facilitate transbilayer move- ments in model systems, namely gel to liquid crystalline phase transitions [578,579], the presence of integral membrane proteins [580,581], the presence of large amounts of phosphatidylethanolamine [582] and, finally, the presence of LPS which is also supposed to disturb the lipid bilayer, since OM-particles and OM-pits, which are visible in

P6

CM

Fig. 8. Molecular organization of the outer membrane of Enter-

obacteriaceae. The most likely positions of outer membrane constituents are schematically indicated. LPS and phospholipid molecules are the major constituents of an asymmetric bilayer. Divalent cations (not indicated) are supposed to play important roles in interactions of LPS. Only three types of proteins have been drawn namely pore proteins including LamB protein (PP), OmpA protein (A) and lipoprotein (LP). Pore proteins have been drawn without interactions with peptidoglycan and lipoprotein, although such interactions cannot be excluded (see subsections IVF-5 and IVF-1, respectively). Several O-antigen chains are much longer than visualized. ECA has not been drawn for reasons of simplicity. Other aspects of the cell envelope like the periplasmic space (PPS) with a nutrient binding protein (BP), the peptidoglycan layer (PG} and the cytoplasmic membrane (CM) with a carrier protein (('P) involved in transport, have also been drawn. For further explanation, see test and Table IV.

freeze-fracture electron microscopic observations of the fracture faces of the outer membrane, are caused by LPS (see Fig. 9, for evidence see subsection IVG-2). However, the latter perturbations are not responsible for the isotropic signal in the ~P- NMR spectra or for any other spectral feature, since the spectra of outer membranes of wild-type cells (rich in particles) of a mutant lacking OmpA protein, OmpF protein and OmpC protein (and therefore deficient in particles) and of LPS-phos-

pholipid liposomes are indistinguishable. This indicates that LPS has probably little diffusional motion around the curved surfaces of the particles, probably due to immobilization by the protein in the particle [574]. In conclusion, the data of X-ray diffraction, freeze-fracture electron microscopy and 3~p-NMR spectroscopy indicate that the outer membrane is largely a planar lipid bilayer with perturbations visible as O'M-particles and (~M-pits.

IVG-2. Nature of O~M particles and OM pits on the fracture faces of the outer membrane observed with freeze-fracture electron microscopy

In the Gram-negative bacteria E. coli [200,308,508,510,570], S. typhimurium [88,571,572], Ps. aeruginosa [511] and Acinetobacter [57~ the outer fracture face of the outer membrane (OM) is densely packed with particles. In E. coli K-12 the particles are 4-8 nm in diameter and occur in estimated numbers of 6000-10 0 0 0 / ~ m 2 [198,508] (Fig. 7A). The opposite inner fracture face of the outer membrane (O~M) contains 3000-5000 pi ts / t~m 2 (Fig. 7B). The latter numbers are probably underestimated [198] and, as the numbers of both particles and pits are decreased in various mutants [~8,200] it is very likely that O~M-particles and OM-pits are complementary [200,570,584]. Around 1974 freeze-fracture particles were generally considered as reflection of membrane proteins. The availability of mutants deficient in outer membrane proteins therefore prompted several investigators to study the freeze-fracture morphology of the outer membrane of E. coli in order to elucidate which proton(s) was (were) involved in the formation of OM-particles [88,198, 200, 308, 508~ 510, 571,572]. In the course of this work it became clear that protein could not be solely responsible [198,200] and that particles with complementary pits must be considered as perturbations of the lipid bilayer by lipids [569,584,585]. By now there is a general agreement that the O~M particles and O~M pits in outer membranes of Gram-negative bacteria are reflections of protein-LPS complexes [2,15,200,511]. It is likely that in the cell protein is required to generate such complexes from which the morphological appearance is determined by the molecular organization of LPS [198,200,363]. In the following we will explain the evidence for this notion in detail as the experiments on the

95

nature of the freeze-fracture particles have been misinterpreted in an important review [2].

Several E. coli K-12 mutants lacking major outer membrane proteins have reduced amounts of both particles and pits especially those lacking two or three proteins. This reduction roughly correlates with the total reduction in amount of major outer membrane protein, although the effect of the OmpA protein is always stronger than that of the OmpF and OmpC proteins [198]. A special problem in these experiments was that the lack of one protein is compensated for by an increase in the amounts of one or more other proteins (Refs. 96, 129, 306 and 586; see also Fig. 6B). Since these compensatory effects are absent in strains lacking all three of these major proteins (Refs. 96 and 306; see also Fig. 6C), the role of various outer membrane proteins in particle formation was studied by comparing mutants lacking the three proteins OmpA, OmpC and OmpF protein with strains which contain large amounts of one of the proteins OmpA, OmpF, OmpC, LamB or PhoE protein [200]. Whereas only about 25% of the surface area of the OM of the former strains was covered with particles (and of the (~M with pits), the presence of each one of the latter five proteins, which resulted roughly in an increase of the protein content of the outer membrane to the wild-type level, lead to an increase to 90-100%. These experiments convincingly showed the capacity of these five proteins in the formation of O~M-par - ticles and OM-pits [200].

The first indication for a crucial role of LPS came from the observation that treatment of E. coli K-12 cells with EDTA, which causes the release of half of the cellular LPS but not of protein, resulted in the loss of about half of the OM-particles thereby causing smooth patches on the OM [198]. These results lead to the hypothesis that LPS complexes form the basis of O'M particles [198]. Reasoning that, if this hypothesis were correct, it might be possible to generate LPS-particles in particle-poor cells, the strain missing OmpA, OmpC and OmpF proteins was incubated with Ca 2+ in the presence of chloramphenicol. Indeed, this Ca 2+ treatment resulted in an increase of the particulated area from 25-30% to 90-100% [200], without affecting the protein or LPS content of the cells. That these newly generated particles and

96

pits, which are morphologically indistinguishable from those in wild type cells, are indeed caused by LPS aggregation, was shown by three types of experiments. Firstly, EDTA treatment of the

~ 4 Ca" -treated cells resulted in particle-poor outer membranes accompanied by the loss of 60% of the cellular LPS without loss of protein. This result stressed the role of LPS by showing that particles and pits with the same morphology as those occurring in the outer membrane of wild-type cells, can be generated by LPS-Ca 2+ complexes. Secondly, when the mutant which lacks three proteins and therefore is particle-poor was replaced by another strain which, in addition, also has shorter, heptose-less, LPS, Ca 2+ treatment did not result in an increase of the number of particles [200]. The most likely explanation for this result is that the LPS-Ca 2 + interactions occur via the heptose-bound phosphate residues of the LPS [200]. Thirdly, purified LPS itself is able to form particles of the same size as those found in whole cells [162].

In conclusion, it has been established that both LPS and at least five different major outer membrane proteins play a role in the generation of particles and pits. The protein component interacts with LPS, resulting in the generation of particles and pits. As the LPS component is responsible for the morphology of these structures, it is likely that only a small portion of the protein is located within the fracture plane [569]. All experimental data are explained by a model in which complexes of various LPS molecules and one or more protein molecules of one protein species are assumed to form a complex, which is visible as an O~M particle with a complementary O~M pit. Thus the population of particles in a wild type outer membrane is a collection of protein-LPS complexes from which in 75-80% of the cases the protein component is one of the proteins OmpA, OmpC or OmpF protein [200]. As LPS without protein can generate such a particle and as EDTA-treated cells with their normal outer membrane protein content have lost about half of the number of particles [198], LPS is likely to play a more direct role in the morphogenesis of the particle than the protein component. The presence of the protein component is assumed to be a prerequisite for LPS aggregation [200]. A molecular model of an (~M particle with complementary OM

pit, which explains all published data, is given in Fig. 9. It should be emphasized that in addition to lipidic intramembranous particles, which have complementary pits and which occur in bacterial outer membranes but also in many other systems including phospholipid model membranes [585], many other intramembranous particles exist which do not have complementary pits and which reflect proteinous structures e.g. those in red blood cell membranes.

O~M particles of Ps. aeruginosa have been studied by Eagon, Gilleland, Stinnett and co- workers. Initially we were unaware of their results, which despite the very different approach, lead to a similar hypothesis. Their results can be summarized as follows. Also in Ps. aeruginosa EDTA treatment results in disappearance of particles. The treatment is accompanied by the release of LPS and, in contrast to E. coli, also of protein. Moreover, small cylinders could be detected in the extract with the diameter of OM particles [511]. The damage to the cells can be restored by incubation in fresh medium and this process requires protein synthesis and energy [587].

The only results containing evidence against our model (Fig. 9) are those of Smit and co-workers [88]. Using LPS mutants of S. t)~phirnurium, they determined the number of particles per ~m 2 and calculated the density of various outer membrane constituents per unit cell surface• They found a perfect correlation between the number of particles and the amount of protein per unit of cell surface [88]. With the present knowledge several comments must be made with respect to their data. (a) The amount of protein was determined with the method described by Lowry et al. [130], which is known to overestimate the amount of protein in some outer membrane mutants [127, 129]. (b). Smi te t al. [88] conclude that the number of LPS molecules per unit of surface area is not influenced by LPS-mutations. Data from other laboratories on similar mutants of E. coli and S. typhimurium indicate that deep-rough mutations result in considerable [94,96] or even dramatic [92,93] increases in the number of LPS molecules• (c). As LPS is now known to form the basis of (~M-particles (Fig. 9), it is dangerous to use LPS mutants for quantifying the number of particles. It is conceivable that such mutations influence fac-

tors like size, number and stability of OM particles.

Some additional comments to the model of O~M particles and OM pits are the following. (i) The presence of units consisting of only one protein species has subsequently been supported by data from stt/dies with chemical cross-linkers (see Table IV). It therefore is likely that the protein component of the particle is a trimer in the case of a pore protein. (ii) As mutants lacking both forms of the lipoprotein have a morphology indistinguishable from that of wild-type cells (Van Alphen, L., Verkleij, A. and Lugtenberg, B., unpublished data) it is likely that the lipoprotein is not involved in the formation or stabilization of particles and pits. (iii) several attempts have been undertaken in our laboratory to study particles and pits in more detail. All biochemical isolation procedures used were unsuccessful. Also attempts to generate two-dimensional crystals by forcing high amounts of only one protein species into the outer membrane, have failed. This was accomplished by either using an ompA mutant of E. coli B E in which over 90% of the remaining outer membrane protein consists of OmpF protein (Fig. 7) or by introduc- ing a multicopy plasmid harbouring the cloned phoE gene in an otherwise particle-poor back- ground (Verkleij, A., Leunissen-Bijvelt, J., Van

Fig. 9. Schematic model of a transversal section through the outer membrane with a LPS-protein complex in the middle, which - upon cleavage of the lip~id bilayer - results in an O'~M particle with corresponding OM pit. Interactions of an oligomeric t ransmembrane protein (in this case a pore protein) with LPS, divalent cations and possibly polyamines are supposed to result in a wedge-shaped organization of the LPS molecules and therefore in the perturbation of the lipid matrix. See also Ref. 200.

97

Boxtel, R., and Lugtenberg, B., unpublished data). Even slow cooling from 37°C to 4°C did not yield regular patterns. (iv) As many proteins involved in the generation of particles are pore proteins, it was hypothesized that most of the particles were the morphological reflections of the intramembranous part of the pores [200]. Indeed, a correlation between the number of particles and the activity of various pores has been reported [363]. (v) In mutants with a decreased number of OM particles it clearly can be shown that both particles and pits are laterally mobile. Slow cooling of the preparations before quenching showed, in addition to large particulate areas, large smooth areas on the O~M [198,200]. In wild-type E. coli K-12 the same effect can be observed except that the smooth areas are much smaller [508]. The reported lack of smooth areas on the O~M of a S. typhimurium Rc mutant [88] is probably caused by quenching from a temperature above the phase transition.

IVG-3. Physical properties of LPS and phospholipids in the outer membrane

The outer membrane undergoes a relatively broad thermotropic order-disorder phase transition over a temperature range of about 20°C as measured with X-ray diffraction [128], 2H-NMR [588-590], differential scanning calorimetry [575,591] and fluorescence spectroscopy [578]. The temperature of this transition range is about 7°C higher than that of the cytoplasmic membrane [590], which can be explained by the higher degree of saturation of the fatty acids of the phospholipids [137] and the lower phosphatidylglycerol content of the outer membrane [590]. These results were obtained using fatty acid auxotrophic mutants fed with various fatty acids. Using wild-type cells of E. coli K-12 the thermotropic phase transition temperature range of the phospholipids appeared to be dependent on the growth temperature of the cells, such that the cytoplasmic membranes are in a mixed gel plus liquid-crystalline state or liquid state at the growth temperature [578]. Wild type E. coli K-12 cells are unable to grow below 8°C, when all the phospholipids are in the gel state [578]. The shift in phase transition temperature range is most likely caused by changes in the fatty acid composition of the phospholipids as a consequence of differences in the growth temperature

98

[ 137]. The transition temperature range of the outer membrane was less sensitive to changes in the growth temperature than that of the cytoplasmic membrane, presumably due to the presence of LPS [592]. The result is a mixed gel plus liquid crystalline state for the outer membrane at all growth temperatures (8 46°C) [578]. X-ray diffraction data indicate that only 25-40% of the outer membrane lipids is participating in the phase transition. This percentage is twice as high in the cytoplasmic membrane [128]. On the other hand, 2H- N M R spectroscopy experiments indicate that all the phospholipids are participating in the order- disorder transition [590]. The phospholipids of the outer membrane certainly are not immobilized [589,590], although they have less motional freedom than those of the cytoplasmic membrane in the phase transition [590]. Restricted motion of E. coli outer membrane lipids has also been reported by Rottem and Leive [593] from electron spin resonance spectroscopy, using spin-labeled fatty acids as a probe. The motional freedom increased strongly when the cells used for membrane isolation had previously been extracted with EDTA, suggesting that LPS plays an important role in the restricted mobility of the lipids in the membrane [593]. Nikaido et al. [542], however, could not detect any restricted motion for the same spin- labeled fatty acids. Spin-labeled fatty acids as probes have the disadvantage that the distribution of the probe over the various lipid domains in the membrane is unknown. Therefore, Takeuchi et al [594,595] used a much more 'natural ' approach by labeling the membranes of E. coli biosynthetically in situ with spin-labeled phospholipids. Spin- labeled phosphatidylglycerol was found to be much more restricted in its motional freedom than phosphatidylethanolamine. The authors explain their result by assuming that phosphatidylglycerol interacts strongly with proteins, while phosphatidylethanolamine probably forms the matrix of the outer membrane. When part of the LPS was removed by EDTA treatment, the mobility of both phospholipids was found to be increased, consistent with the data of Rottem and Leive [593] with spin-labeled fatty acids. All these experiments, except those of Nikaido et al. [542], indicate that the outer membrane phospholipids are more immobilized than those of the cytoplasmic

membrane. However, only Nikaido et al. used outer membranes still connected to the peptidoglycan layer while the other data [593] were obtained with outer membranes from spheroplasts. The latter isolation procedure is known to result in randomization of LPS, and therefore probably also of phospholipid, over both monolayers of the outer membrane [89]. Such a randomization could explain the strong influence of LPS on the motional freedom of the phospholipids. It would be interesting to repeat the experiments of Takeuchi et al. [594,595] with outer membranes connected to peptidoglycan.

LPS is not only able to participate in phase transition, but shows a phase transition itself. X-ray diffraction [577,592]~ light scattering [170] and a di fferential scanning calorimetry [ 170] experiments with E. coli LPS revealed a rather broad transition [170,577] in which the midpoint temperature and the transition range were dependent on the growth temperature of the cells from which LPS was isolated [170]. A lower growth temperature was correlated with the occurrence of more unsaturated fatty acids and a lower transition temperature in light scattering, the disappearance of a transition observed with X-ray diffraction, and an increase in the fluidity of lipid A as measured with electron spin resonance spectroscopy [ 170,592,596]. Using freeze-fracture electron microscopy on purified LPS several transitions were observed between 22 and 37°C at temperatures which were strongly dependent on the hydration conditions and the degree of purity of the LPS preparation [144,162,597]. The broad range of transition might be explained by the heterogeneity in the composition of LPS, both with respect to the various substituents [165,599] and to the length of the sugar chain [ 142,183-185,597,598].

The diffusion constant of the phospholipids in the outer membrane as determined with ESR spectroscopy is as high as in the cytoplasmic membrane and in model membranes (D = 2.5-10 s cm2/s) [542], implying that a phospholipid molecule moves over the length of the E. coli cell in 1 s [138]. The diffusion constant for LPS is about 3 • 10 13 cm2/s [600], determined by following the rate of distribution of newly synthesized LPS over the cell surface. In these experiments a galE mutant of S. (vphimurium was used and a pulse of corn-

99

plete LPS was synthesized by shifting the cells from galactose-less to galactose-containing medium. This LPS was detected using ferritin-labeled antibodies specific for the O-antigen [600]. With similar experiments it was shown that LPS diffuses from its 50 sites of insertion in the outer membrane over the surface of the cell [35,600,601] into domains [537]. This LPS can be isolated and does not mix with 'old' LPS for at least one generation [537]. In E. coli and S. typhimurium two classes of LPS have been recognized, one of which is extrac- table with EDTA [85]. The latter LPS fraction is not identical to the fraction of newly synthesized LPS described above, since extraction of cells with EDTA [85] resulted in extraction of both newly synthesized and 'old' LPS [537].

Lateral diffusion constants of phopsholipids, LPS and pore proteins were determined very elegantly with the 'fluorescence redistribution after photobleaching' technique by Schindler et al. [602,603]. They found that the diffusion coefficient for both LPS and phospholipids in LPS-phospholipid (1 : 1, w/w) liposomes is about 10 - 9 c m 2

• s ~ at 24°C. For phospholipids this value is an order of magnitude lower than in pure phospholipid liposomes. The mobility of the phospholipids is hardly influenced by the incorporation of pore protein in these mixed bilayers, whereas the mobility of LPS decreases ten-fold when liposomes containing 60% protein were used. The diffusion constant for the protein was < 10 12 cm 2 . s- ~. After fusion of cells with liposomes containing spin- labeled LPS, Schindler et al. determined the diffusion coefficient of LPS in outer membranes as 2 . 1 0 l0 cm 2" s - l , indicating more restricted motion than in liposomes. This value is much higher than that obtained by M~ihlradt et al. [600]•

IVG -It. Interactions of proteins with the lipid matrix (see also Table IV)

To our knowledge the first indications for LPS-protein interactions for E. coli, at least in vitro, were published in the early seventies [604,605]. The first indications for in vivo interactions came from analyses of the proteins and LPS in mutants with incomplete structures. In deep- rough mutants of E. coli [87,93,94,96,236] and S. typhimurium [88,90,92] decreased amounts of outer membrane proteins, especially of OmpF protein,

PhoE protein and protein III were found. The existence of interactions of LPS with the proteins OmpA, OmpC, OmpF, Tsx, LamB and PhoE protein was clearly established by using the property that LPS was required for in vivo a n d / o r in vitro phage receptor activity ([ 171,217,236, 237,293,422, 526]; see also Table IV). Additional evidence for an interaction of pore proteins with LPS comes from the observation that the presence of LPS stimulates the binding of these proteins to peptidoglycan [522,606]. Moreover, the presence of LPS is required for in vitro pore activity of OmpF protein [362]. In the case of OmpA protein, several properties like resistance against protease and heat denaturation [526], electrophoretic mobility, receptor activity for phages and for donor cells in conjugation [217,308,381] are dependent on the interaction with LPS. Lipid A is essential for these properties, although the core sugars may also play a role as the rate of adsorption of phage K3 to vesicles containing lipid A and OmpA protein is strongly reduced compared to the rate measured in the same system containing complete LPS [171]. Freeze-fracture electron microscopic studies strongly suggest that the majority of the particles on the outer fracture face with corresponding pits on the inner fracture face consist of complexes of LPS with either OmpA, OmpC, OmpF, PhoE or Lamb protein (see subsection IVG-2).

In order to obtain a better understanding of the interaction between OmpA protein and LPS, the interaction of phage K3 both with its receptor in whole cells and with its reconstituted receptor was examined in detail [170]. The temperature dependence of the adsorption rate of phage K3 to OmpA protein-LPS complexes (containing LPS isolated from cells grown at either 12°C or 37°C) is similar to that of the adsorption rate of the phage to the corresponding cells, showing that the reconstituted complex imitates the in vivo situation very well. Therefore it was concluded that in wild type cells OmpA protein-LPS complexes do indeed occur. In an Arrhenius plot of the rate of phage adsorption to the OmpA protein-LPS complexes, two inflection points were detected [170] at temperatures which fall in the transition range of the outer membrane (subsection IVG-3). The inflection point temperatures are dependent on the fatty acid compositions of LPS [170]. Moreover, since the higher

lOt)

inflection point temperature, both for LPS from cells grown at 12°C and at 37°C, was also found in pure LPS in both light scattering and differential scanning calorimetry measurements, it was concluded that this transition is a thermal one. ~ P-NMR measurements showed that at the higher transition temperature LPS is more mobile [162]. This increase in fluidity could be correlated with differences in the fatty acid composition of LPS isolated from cells grown at 37°C and at 12°C [170]. A similar increase in the fluidity of LPS was observed in electron spin resonance spectroscopy when lipid A from Pr. mirabilis cells grown at 43°C was compared with that of cells grown at 15°C [596]. The transition of purified LPS from small bilayer ribbons to large vesicles, observed by freeze-fracture electron microscoppy [162], might be related to this thermal phase transition. Since the lower inflection point temperature could not be correlated with a transition in isolated LPS, but as it is influenced by the type of LPS which is incorporated in the protein-LPS complexes, it is likely that the interaction between OmpA protein and LPS is involved in this transition. This transition may be detected with ESR, using spin-labeled LPS or with 2H-NMR measurements, using de- uterated LPS. In order to investigate whether the phase transitions found for the OmpA protein-LPS complexes generally occur in protein-LPS complexes, it would be interesting, but extremely tedious, to study the phase transition characteristics of other phage adsorption processes, in which outer membrane proteins are involved.

The phase transition behaviour of the OmpA protein-LPS complexes was strongly influenced by the presence of phospholipids. The phase transition characteristics of the inactivation of phage K3 by OmpA protein-LPS-phospholipids differed from those of OmpA protein-LPS complexes and the characteristics of the adsorption of phage K3 to cells of a mutant containing wild-type LPS, but increased amounts of phospholipids in its outer membrane, part of which is present in the outer monolayer (see Fig. 6B), differed from the phase transition characteristics of the adsorption to wild-type cells in a similar way [170]. These results indicate that in wild-type cells phospholipids do not influence the receptor domain of OmpA protein-LPS complexes [170], a conclusion consistent

with a model in which wild-type cells harbour no phospholipid in the outer monolayer of their outer membrane (Fig. 8).

IVG-5. Distribution of outer membrane constituents over both monolayers

Nikaido and co-workers were the first who calculated the distribution of the various outer membrane constituents over both monolayers [88,607]. These calculations are based on the knowledge of the surface covered by the individual molecules, of the amounts of the various outer membrane constituents relative to the mass of the cell and of the surface to mass ratio of the cells [88]. However, the surface of the cell is very difficult to determine since in each culture cells of various sizes occur, and since artefacts may be introduced during the preparation of samples for electron microscopic observation. The surface area occupied by phospholipids and lipopolysaccharide can be estimated from measurements in model systems, but this is impossible for proteins for which the tertiary structure is unknown. Smit et al. [88] therefore determined the surface occupied by the proteins by subtracting the surface of LPS and phospholipids from the measured total cell surface.

In order to circumvent the surface measurements we have used another approach to determine the distribution of outer membrane constituents over both monolayers. It is based on the observation that the lack of the proteins OmpA, OmpC and OmpF protein in E. coli K-12 strain P692tut2dl is compensated for by increased amounts of phospholipid and LPS [127]. We therefore reasoned that the bilayer space occupied by these three proteins in the parent strain P400 is equal to that occupied by the increased amounts of phospholipids and LPS in the mutant strain P692tut2dI. Furthermore, as these three proteins together with the lipoprotein account for 95% of total outer membrane protein (see legend of Table V), we assumed that the bilayer surface occupied by the total outer membrane proteins was equal to this area plus the area occupied by the lipidic part of the lipoprotein, probably resulting in a small underestimation of the space occupied by the outer membrane proteins. The results of our calculations are summarized in Table V. The surface area occupied by a lipoprotein molecule

101

can be estimated as about 0.8 nm 2 since according to the models of McLachlan [552] and Braun [266] only the fatty acids of the lipoprotein are embedded in the bilayer region and since these fatty acids are linked to a glycerol residue as in tri- glycerides. This results in an almost negligible contribution of the lipoprotein to the bilayer area (see footnote g of Table V). The surface area occupied by a phospholipid molecule is 0.59 nm 2 as determined with monolayer experiments on total E. coli phospholipid at maximal surface pressure [608]. This is in good agreement with the value found for S. typhimurium [176]. From recent de- terminations of the surface area of a monomer of LPS in monolayer experiments a value of 2.50 nm 2 can be deduced [139]. The results of these calculations (Table V) indicate that about one half of the surface area of the inner monolayer and about one third of surface of the outer monolayer is occupied by protein, whereas phospholipids and LPS, respectively, cover the remainder of these mono-

layers. Smit et al. [88] calculated that approxi- mately half of the outer monolayer is occupied with lipopolysaccharide and three quarter of the inner monolayer with phospholipids. The differences between their values and ours cannot be ascribed to inaccuracies in the surface determina- tions alone. Other factors may be (i) most recent value for the surface area occupied by a LPS molecule is three times higher than the value determined previously [176], and (ii) differences in protein : LPS : phospholipid ratio, probably caused by an overestimation of the amount of protein as a consequence of the use of the method developed by Lowry et a1.[127,129] and by differences in growth conditions and strains (compare subsection IIIA).

IVH. Molecular organization of the outer membrane of Enterobacteriaceae

Based on the data of the previous sections of this chapter, a model (Fig. 8) can be constructed

TABLE V

C A L C U L A T E D CONTRIBUTIONS OF PHOSPHOLIPID, LPS A N D PROTEINS TO THE SURFACE AREA OF THE OUTER M E M B R A N E OF ESCHERICH1A COLI K-12 STRAIN P400

Component Composition Average Molar Surface area (wt%) a molecular ratio per molecule

weight (nm 2 )

Fraction of surface area in % g

Outer Inner monolayer monolayer

Phospholipid 19.5 700 27.9 0.59 c 0 46 LPS 43.7 4500 b 9.7 2.50 d 68 0 Major protein

(OmpF, C, A) 19.8 36000 0.55 _ e 32 50 Lipoprotein 15.2 7 200 2.1 0.8 f 0 4

a Derived from Ref. 127.

b From unpublished observations and data from Ref. 181. LPS is considered to be a monomer [165]. c From monolayer studies by Haest et al. [608] on total phospholipid of E. coli. d From monolayer studies by Fried and Rothfield [139] on LPS of S. typhimurium strain G30(galE), which has a structure similar to

that of E. coli K-12.

c The surface area occupied by individual protein molecules is not known. The amount of bilayer space occupied by OmpA, OmpC and OmpF proteins in E. coli K-12 wild-type cells was calculated by assuming that the bilayer space which is not occupied by these proteins in cells of the OmpA, OmpC and OmpF protein deficient strain P692tut2dl is filled with LPS and phospholipid.

f Only the fatty acids are assumed to occupy space in the bilayer [552], predicted to correspond with 1.5 times the surface of a phospholipid molecule (see text).

g The outer membrane of strain P400 is considered to be asymmetric, in that LPS is exclusively located in the outer monolayer and phospholipid and the lipoprotein exclusively in the inner monolayer (see text). The sum of the amounts of OmpA, OmpC and OmpF protein and the lipoprotein is assumed to be equal to total outer membrane protein for the calculations of the surface areas occupied by the proteins (c.f. note e), since the total amount of OmpA, OmpC and OmpF protein is 54% and the amount of the lipoprotein is 40% of the total outer membrane protein. The latter value is obtained from the weight ratio of the sum of OmpA, OmpC and OmpF protein (about 2.105 copies per cell with an average molecule weight of 36000) over total lipoprotein (about 7.5.103 copies per cell, molecular weight 7200).

102

for the outer membrane of wild-type cells of E. coli and probably also for thai of other Enterob- acteriaceae. Only the pore proteins, OmpA protein, lipoprotein, LPS and phospholipids are included in this picture, since only their localization and interactions have been established reasonably well. The most important properties of the outer membrane will be discussed here using this model.

1. Diffusion of hydrophilic solutes occurs via general pores which consist basically of trimers of pore proteins complexed with LPS. The specificity of the pore is determined more or less by the properties of the pore protein and, at least in the case of the LamB protein, by the periplasmic maltose-binding protein.

2. The outer membrane is hardly or not at all permeable for hydrophobic substances. Permea- tion of hydrophobic substances between LPS molecules is unlikely since these molecules are strongly negatively charged in the core-lipid A region. The LPS molecules are held together by positively charged divalent ligands (especially Ca 2+) [158,197,199], thereby complexing LPS in various ways [158,162]. The impermeability of the outer membrane for hydrophobic substances has been attributed to the absence of phospholipid bilayer regions in the outer membrane [361]. In- deed it has been shown that small hydrophobic molecules dissolve in a lipid bilayer [609,610]. We have already described that pore protein deficient mutants have increased amounts of phospholipid, part of which is located in the outer monolayer. This is also considered to be the explanation for the sensitivity of these mutant cells for SDS [236]. A similar effect may be caused by the treatment of P. mirabilis cells with the antibiotic cerulenin in the presence of exogenous fatty acids. Such a treatment makes the cells sensitive to the antibiotics vancomycin and rifampicin and results in a decrease of the LPS content by 30-50% [611]. Although the outer monolayer is certainly poor in phospholipids it may be an exaggeration to say that they are absent in that monolayer. Nixdorff et al. [163] have shown that the presence of LPS protects phospholipids in liposomes against solubilization by detergents. Therefore such shielding effects might have prevented the detection of small phospholipid-containing areas in the outer leaflet. With the presence of such areas processes like

fusion with liposomes [136,140] and blebbing [47,272] are easier to understand.

3. The outer membrane of an E. coli K-12 mutant in the envA gene, contains a decreased amount of LPS and has an increased influx of hydrophobic drugs. This phenotype could be suppressed by a second mutation which brings about an increase in the amount of outer membrane protein [612]. Considering the model of Fig. 8, we favour the following explanation. The decreased amount of LPS due to the envA mutation is partly replaced by phospholipid. The resulting phospholipid bilayer explains the sensitivity to the antibiotics. This phenotype can be suppressed by a second mutation, sefA, which causes an increase of the protein content a n d / o r a decrease in the phospholipid content resulting in the virtual disappearance of phospholipid from the outer monolayer. In the light of the often occurring protein- LPS interactions it should be noted that an increase in outer membrane protein content, accompanied by a still lower LPS content [612], seems only possible if not all LPS present was already involved in LPS-protein interactions. In other words, the results are consistent with the presence of substantial amounts of LPS in interactions which differ from protein-LPS interactions. LPS- LPS interactions seem to be the only possible alternative and this explanation is consistent with both the model of Fig. 8 and with the observation that LPS is present in various domains [537].

4. During bacteriophage infection and bacterial conjugation specific processes, of which most details are still unknown, are involved in the penetration of large DNA molecules into the cell. Some phages push their injection needle into the cell after recognition of the receptor, likely in a mecha- nical way [613]. Since zones of adhesion are likely involved, non-bilayer conformations of the membrane might be important for this process (see Fig. 2).

Naked DNA can be brought into E. coli cells in a very inefficient way by a process called transformation. This can only occur in competent cells, i.e. cells which have been treated with high concentrations of Ca 2+ and which subsequently have been given a temperature shock [614,615]. The two components of this treatment, the effect of Ca z~ and the passage through the phase transition, are

103

known to promote non-bilayer organization of lipids. DNA thus may enter, in a very inefficient way, through such temporary perturbations in the membrane. A decrease of the outer membrane barrier can also be induced by Ca 2÷ treatment alone [616]. This causes a less drastic effect as it is not sufficient for obtaining transformation-competent cells (Van Die, I.M., Bergmans, J.E.N. and Hoekstra, W.P.M., unpublished data).

5. The outer membrane is a dynamic structure, in which newly synthesized material is inserted and from which certain parts are excreted as blebs. Most of these mechanisms are unknown. Essential is that the structure allows such changes. It seems likely that non-bilayer phases in zones of adhesion play an important role (see Fig. 2). Moreover, since experiments in which vesicles have been fused with cells indicate that the lipids are exchanged between the two membranes [136,140], a similar process might be involved in the biogenesis of the outer membrane.

1VI. Outer membrane of other Gram-negative bacteria

Very little is known about the structure of the outer membrane of other Gram-negative bacteria. The occurrence of proteins similar to lipoprotein, OmpA protein and pore proteins has been described in various species. Since several Gram- negatives like Neisseriaceae and H. influenzae are very sensitive to hydrophobic antibiotics, it seems reasonable to predict that their outer membranes contain phospholipid bilayer regions. Indeed, in a recent paper evidence for this idea is presented in the case of N. gonorrhoeae [617]. The presence of such phospholipid bilayer areas can be a reason why certain Gram-negative bacteria shed off numerous outer membrane blebs. A low content of those outer membrane proteins involved in anchoring the membrane to,the peptidoglycan layer is also likely to contribute to this phenomenon.

V. Future prospects

The knowledge of the architecture and function of the outer membrane of E. coli and S. typhimurium has reached an advanced state, although many questions are still unanswered. For

the near future progress can be expected in our knowledge of the standard laboratory strains as well as in that of strains which are more relevant to men, animals or plants.

Since it is clear that LPS is a heterogeneous molecular species (see subsection IIIC-3) there is a need for application of better separation methods, for instance high pressure liquid chromatography. The significance of the separated species for the various functions of lipopolysaccharide can subsequently be investigated in vitro. The physico- chemical behaviour of LPS is largely unknown [ 162]. Extensive NMR-studies (both 31P-NMR and 2H-NMR) will be required on complete LPS as well as on partial degradation products in order to understand its complex properties as a membrane component.

Detailed information on the structure of the pore proteins will be available when crystals of the purified proteins are large enough for crystallographic analysis of the tertiary and quaternary structure of the pore proteins [373]. The structure- function relationship of outer membrane proteins will be studied using chemically modified proteins. Site-specific mutations in cloned DNA will be used to bring about any desired mutation. Also the use of monoclonal antibodies will be a great help in elucidating the topology of surface-exposed sites. Furthermore, the availability of cloned DNA will enable us to study the regulation of the synthesis of outer membrane constituents in detail. It seems likely that many details of the exiting area of biogenesis of outer membrane proteins will be known within a few years as cloning and site- specific mutation enable us to study the role of the signal peptide and of other important features of the molecule.

Gram-negative bacteria occur in a wide variety of environmental circumstances. For example, Chlamydia lives intracellular [618], Pseudomonas in surface water and soil, E. coli in the digestive tract, N. meningitidis in the upper respiratory tract and Agrobacterium in plants. This implies that these organisms have totally different requirements for growth and that their outer membranes meet various environmental conditions. Comparison of membranes of various organisms has just started. Moreover, within certain bacterial species large differences have been observed. For example E.

104

coli is a harmless inhibitant of the intestine of many animal species, but some strains are pathogenic causing severe diarrhea whereas other strains can cause urinary tract infections, they are able to invade the host organism and to grow in the blood stream of in the cerebrospinal fluid causing the very severe infectious diseases sepsis and meningi- tis. Complex mechanisms are involved in adherence, invasion of the tissue, and in coping with the many defense mechanisms of the host ]17]. Surface structures of the cell are involved and data are beginning to appear showing that fimbriae, outer membrane proteins, capsular polysaccharides and LPS are involved in several aspects. Mechanisms like antigenic variation, immunosup- pression, molecular mimicry of host components and toleragenicity are major mechanisms used by bacteria [619]. An enormous field of research is open to elucidate these mechanisms, in which outer membranes certainly play a role [619]. For the development of vaccines (Refs. 534, 535 and 620, and Dankert J., Hofstra, H. and Veninga, T.S. (1980) in FEMS Symposium on Microbial En- velopes, Saimaanranka, Finland, Abstract 51) and diagnostics [621] important features like surface localization and immunogenici ty have to be determined. It also has to be established whether antigenic determinants are common or specific. This field is rapidly developing and vaccines for Neisseriaceae and H. influenzae are already in field trial [622-626]. Impor tan t combinat ions of biochemical and immunological techniques have recently been developed [531,627-629] including the construct ion of cell lines producing monoclonal antibodies [630,631]. Molecular genetics can be used to clone D N A coding for important constituents in E. coli K-12. The cloning of genes for E. coli K1 capsule formation is a recent example [632]. The recently constructed colony bank of cloned D N A of virulent Treponema pallidum [633] might enable the investigator to express and produce antigens of this important pathogen in E. coli K-12. As T. pallidum cannot be grown on laboratory media, such an approach would be the only possibility to develop a vaccine. However, troubles might arise in the expression of foreign outer membrane constituents in E. coli K-12, e.g. because of the possibly strict requirements of protein-LPS interactions [304,393,535,634]. On the

other hand, it has been shown that expression of O m p A protein of various Gram-negative bacteria into E. coli did not meet serious problems [315]. It can also be expected that the near future will be important for the development of peptide vaccines [632]. The second decade of outer membrane studies will be as competative and laborious as the first one and certainly even more exciting.

Acknowledgements

The authors greatly enjoyed the many fruitful discussions with Arie Verkleij. They thank V. Braun, S.T. Cole, C.F. Earhart, R.E.W. Hancock, U. Henning, M. Inouye, P.M. M~ikel~i, S. Mizushima, T. Nakae, J.B. Neilands, H. Nikaido, J.P. Rosenbusch, C.A. Schnaitman, B. Withoh and H.C. Wu for providing them with unpublished results. We thank Lia Claessens, Winny Geelen and Joy Zantkuyl for typing the manuscript and Dick Smit for excellent art work.

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