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The morphologic structure of isolated bacterial lipopolysaccharide
J. Mol. Biol. (1967) 25, 15-21
The Morphologic Structure of Isolated Bacterial Lipopolysaccharide J. W. Department
SHANDS,
J. A. GRAHAM
JR.,
of Microbiology,
University
Gainesville,
AND
K. NATH
of Florida College of ilfedicine
Florida,
U.S.A.
(Received 17 October, 1966) The morphology of lipopolysaccharides isolated from smooth and rough strains of Salmonella typhimurium was studied by electron microscopy using a positivestaining procedure. When viewed face on, lipopolysaccharide from the smooth strain was a homogeneously stained ribbon which could be broken down to homogeneously stained discs by sonication or agitation with ether. When viewed edge on, the ribbon and disc had a trilaminar appearance reminiscent of cross-sections of arrays of bimolecular leaflets. Lipopolysaccharide from a mutant lacking Opolysaccharide side-chains had a similar appearance and identical dimensions, whereas that from a heptoseless mutant, lacking 0 and “core” polysaccharide had smaller and more delicate trilaminar structures. The behavior of lipopolysaccharides in lipid solvent and the common morphological feature found in lipopolysaccharide regardless of the polysaccharide content suggest that the major morphological determinant is the lipid moiety. There is suggestive evidence that the “core” polysaccharide contributes to the trilaminar structures, whereas the 0-polysaccharide side-chains do not.
1. Introduction The lipopolysaccharides of Gram-negative bacteria are macromolecular complexes which possess both endotoxic and 0-antigenic activities. Chemically these substances are composed of a lipid moiety (lipid A) and a polysaccharide moiety, the proportions of which differ from one species to another and even among different strains of the same species. Recent investigations of LPSt f rom Salmonella (Nikaido, Naide $ MLkelS, 1966; Liideritz et al., 1966; Osborn, 1966)indicatethatthelipopolysaccharides are composed of repeating sugar units which impart the antigenic specificity. These are attached to a polysaccharide “core” possessing a “backbone” of heptose phosphate and 2-keto-3-deoxyoctonate, which in turn is linked to lipid A. Morphologically, in electron micrographs of shadowed specimens, LPS from Escherichia coli has the appearance of 100 A droplets at neutral pH or long ribbons at pH 10 (Schramm, Westphal & Liideritz, 1952). Similar morphology was demonstrated in LPS from a strain of E. coli (Weidel, Frank & Martin, 1960) and in that from Bordetella pertussis (Milner et al., 1963). In a detailed study of Boivin-type LPS from E. coli Beer, Braude & Brinton (1966) found a variety of shapes: snake-like, donut-like, flat sheets and slender rod-like filaments. The latter particles possessed the t Abbreviation
used: LPS. lipopolyaaccharidp. 15
IA
J. IV.
SHANDS,
all..
J. 11. (:I<.-\H;\N
ASD
I<. SAi’I’H
greatest toxicity, indicating t,hat, t,hey. might, bc responsible for the tosicif,y of ihe crude extract. Attempts to correlate the biochemical and morphological data ha.ve led t’o t’ht: proposal of two possible models. On one hand, Milner et rtl. (1963) suggested that LPS is a cellulose-like “micellar structure” composed of aggregated bundles of haptenic polysaccharide. On the other hand, Rothfield, Takeshita, Pearlman & Horne (1966) proposed that LPS forms a phospholipid-like, ordered “leaflet structure”. This paper is concerned with additional observations of the morphology of LPS in an attempt to correlate morphology and chemical structure. Advantage is taken of the afEnity of LPS for certain heavy-metal stains which delineate structure not seen in specimens prepared by the shadow-cast technique. In addition, the recent availability of strains of Gram-negative bacteria deficient in one or more parts of their lipopolysaccharide now allows one to correlate specific structural changes with chemical differences. Purified LPS from a wild strain of Salmonella typhimurium was compared with those of two mutant strains, one lacking the 0-polysaccharide side-chains, the other lacking both side-chains and “core” polysaccharide.
2. Materials and Methods (a) Bacteriu A smooth strain of Salmonella ty$Gmurium (strain 7) was obtained from Dr M. Herzberg (Dept. of Bacteriology, University of Florida). The surface characteristics and cell wall composition of this bacterium have been described (Herzberg & Green, 1964). The mutants of S. typhimurium LT2, kindly provided by Dr B. A. D. Stocker (Dept. of Microbiology, Stanford University), were (1) TV1 19, an RI1 mutant, which possesses a lipopolysaccharide devoid of specific “0” side-chains, and (2) SL1102, a variant, the LPS of which contains no heptose. The lipopolysaccharide of this latter strain is composed of lipid A and &-k&o3-deoxyoctonate, the latter being the only carbohydrate component detected (Stocker, Wilkinson & M&e%, 1966). E. co&i 0113 was obtained from Dr E. Ribi (Rocky Mountain Laboratories, Hamilton, Montana). The smooth strains of S. typhimurium and E. coli were cultivated in glucose-mineral salts medium for 18 hr at 37’C on a shaker while the mutants were grown in Brain Heart Infusion Broth (D&o) under the same conditions. The bacteria were harvested by cent*rifugation and washed twice with distilled water. (b) Preparation
of cell walk
After washing, the bacteria were suspended in a small volume of water and disrupted in a French pressure cell at 14,000 lb/sq. in. The broken and intact cells were separated from the protoplasm by centrifugation at 27,000 g for 15 min. The pellet was resuspended in O-1 M-phosphate buffer (pH 8-O), and 10 mg of powdered trypsin (Difco) was added for each ml. of pellet. Following incubation with gentle stirring for 10 hr at 37”C, the walls were washed repeatedly in 1.0 M-N8c1 and in distilled water. Walls were separated from intact bacteria by gently resuspending them and by discarding the dense button which formed at the bottom of the tube during centrifugation. Washing was continued until the cell walls were judged clean by electron microscopy.
(c) Extraction
of lipopolysaccharide
LPS was extracted from cell walls by the procedure of Westphal, L;ideritz dt Bister (1952). Approximately 600 mg of cell walls were suspended in 100 ml. of distilled water and an equal volume of liquid phenol was added. The mixture was stirred for 30 min at 60°C following which the phenol and water phases were separated by chilling and centrifugation, and the water phase was removed. The procedure was repeated after the addition of 100 ml. of water to the phenol phase. The two water phases were combined, dialyzed in dist)illed water and lyophilized.
PLATE III. x 330,000.
LPS
from
8. typhimuriwn
strain
i stwind
with
umnyl
acetate.
x 100.000:
inxct,
PLATE XIV. Diagram showing possible intwpretati?n of’ various image+ ohsrrverl in micrographs in terms of bimolecular leaflet structure. A. Flat disc (observed after sonic&ion or ether treatment) is a trilaminar structure vicwcd face on. 8. Occasional donut-shaped part,irle may rnprrsent trilaminar structure with ~~rt~y~al faces. (‘. Twisted ribbon strurt,urr.
LIPOPOLPSdCCHARIDR
~IORPHOLOGT
17
(d) Electron microscopy Lipopolysaccharide was suspended in glass-distilled water at a concentration of 1 mg/ml. LPS extracted from S. typhimurium 7 was stained positively by adding a drop of 2% in-any1 acetate to 0.3 ml. of the suspension. A drop of this mixture was placed on a carboncoated grid and then quickly removed by touching filter paper to the edge of the drop. Since this procedure precipitated LPS derived from the rough strains, these materials were stained by inversion of a grid containing dried LPS onto a drop of many1 acetate for 1 I,O 8 min, followed by gentle washing with water. Shadow-casting was carried out with uranium oxide in an Hitachi HUS-313 vacuum evaporator. Sectioned LPS from strain 7 was prepared by precipitation with ferritin-labeled antibody, the preparation and specificity of which have been described (Shands, 1965). The precipitate was washed 3 times in 0.1 M-phosphate buffer (pH 7.2) and fixed for 1 hr in 1 ‘J; osmic acid using the same buffer. The precipitate was then pelleted in 1% bovine serum albumin in a small polyethylene tube, the supernatant fluid was aspirated from the pellet, and a drop of 25% glutaraldehyde was added to coagulate the protein. The coagulum of LPS in albumin was cut into small blocks, dehydrated in alcohol, and embedded in Epon 812. Thin sections were cut with the Porter-Blum MT-2 microtome (Ivan Sorvall, Norwalk, Conn.) and stained with uranyl acetate. The materials were examined with a Siemens Ehniskop I electron microscope. ,Most pictures were taken at a plate magnification of 40,000. Dimensions were determined by means of a Leitz microscope equipped with a micrometer eye-piece.
3. Results Plate I shows positively-stainedlipopolysaccharide extracted from S. typhimurium 7. The appearance is that of a flat ribbon which is primarily linear but which branches freely. Its diameter varies but averages 160 A. At infrequent intervals the ribbon narrows to 90 A, and in such places (indicated by arrows) there is a triple-layered structure which consists of two dense outer lines enclosing a less dense cent’er. The general morphology depicted in Plate I is consistent with that demonstrated by others with shadow-casting. However, as additional evidence that the large structures stained are LPS and that others are not missed by the staining procedure, the same preparation was stained with uranyl acetate and then shadowed lightly with uranium oxide. The result is represented in Plate II, which shows that shadow-casting does not expose any macromolecular structures which are not also stained. The trilaminar structure revealed in Plate I raises at least two possibilities. Since its appearance is infrequent, it is possible that it is introduced into the LPS at different points. An alternate possibility is that the entire ribbon is trilaminar, but because of asymmetry tends to rest on its widest surface. The trilaminar appearance would, therefore, be visible only at twists in the ribbon or when the ribbon was made to stand on its thin edge. Several observations indicate that this latter possibility is the correct one. Plate III shows stained LPS and an enlargement of a twist. A distinct trilaminar structure travels from one edge of the flat ribbon to the opposite edge. Plates IV through VII show the effects of dispersing LPS. The material in Plate IV was dispersed by sonication in a Branson Sonifier at 20 kc/s at maximum power for 3 to 5 minutes. It is evident that the long ribbons have been broken up into smaller particles. Some of these are flat, homogeneously stained discs with diameters up to 500 A (A), whereas many other particles (B) have a trilaminar appearance. The latter probably represent flat discs standing on edge, since some particles seem to be resting at an angle and have both trilaminar and disc appearance (C).
IS
J. IV.
SHANDS,
JR..
J. A. C;RA IHAM
ASI)
K.
S.i’l‘I-I
Plate V shows LPS dispersed with 0.5% Tween 80 for 10 minutes at 20°C and stained in the usual manner. Two effects are noteworthy. First, t’he long ribbons arc broken into shorter segments; and second, t,he width of the ribbon is reduced from 160 to 116 8. The width and thickness, therefore, approximate one another: namely. 116 x x 94 .&. With these changes, one might expect, to see more trilnminarstructure, as iti evident in Plate V. =\fter the dispersion in Tween 80, LPS is often trapped in droplets of Tween. Plate VI illustrates this situation and demonstrates that the dimensions of the LPS trilaminar structures may vary. In some areas they appear compressed, and in others they appear expanded. Plate VII shows a preparation of LPS dispersed by shaking it vigorously for several minutes in water saturated with diethyl ether. The result is quite similar to that of sonic dispersion. One finds flat discs (A) and occasional donut-sha,ped particles (C). In thicker areas of the preparation, there are numerous trilaminar structures and occasional particles which seem to be resting at an angle which appear as trilaminar st*ructures and discs (B). Jf LPS is constructed entirely as a trilaminar structure, and if this appears infrequently because LPS tends to lie on its widest surface, then in sections of LPS, where the orientation is random, one should see much more trilaminar structure. Plate VIII shows a lipopolysaccharide preparation which has been precipitated with ferritin-conjugated antibody, embedded, sectioned and stained. The amount of trilaminar structure visible is obviously increased. The structures demonstrated so far are from a phenol-water extract of cell walls of S. typhimurium 7. To rule out the possibilities that the structures are artifacts proa duced by the extraction procedure or are found only in extracts of Xalmonellu, milder extraction procedure was used on whole cells of S. typhimurium 7 and E. coli 0113, namely, the aqueous ether method of Ribi, Milner & Perrine (1959). Plates IX and X show the LPS extracted from E. coli and X. typhimurium, respectively. Although neither of the preparations appears morphologically as well defined as the phenol ext)ract of cell walls, similarities are easily identified. Both of these preparations contain trilaminar structures and one contains the familiar ribbon. The amorphous appearance of the remainder of the materials may be due t,o contaminants or t,o the higher protein content of aqueous ether extracts which ma,y alter the morphology. Lipopolysaccharide extracted from rough mutants of S. typhimurium which lack the 0-polysaccharide side-chains and both side-chains and “core” polysaccharide were also found to contain trilaminar struct#ures. Plate XI shows a stained sample of LPS from the RI1 mutant TV119. Although the material tended to clump, many triplelayered structures are visible. In spite of the fact that this LPS possesses no Opolysaccharide side-chains (Stocker, et al. 1966), its dimensions are the same as those of t’he wild strain 7 LPS. Plate XII shows LPS derived from strain SL1102 which lacks both side-chains and “core” polysaccharide. Like the other preparations, it had been lyophilized and reconstituted in distilled water. Peculiarly, its morphology is unlike that of the other preparations, since it has an appearance which suggests folded and indented vesicles. This appears, however. to have been the result of lyophilization, since a sample of the same preparation which had not been lyophilized produced many triple-layered structures (Plate XIII). A comparison of these trilaminar structures with those in LPS of the wild strain and RI1 mutant shows that they
LIPOPOL~~ACCHAI~IUE
AIoL~L’HOI,C)GY
I9
arc more delicate, having less dense outer lines and a smaller over-all dimension, namely, 60 A versus 90 A.
4. Discussion The illustrations show that lipopolysaccharide derived from a smooth strain of &. typhimurium by the phenol-water procedure is a ribbon-like structure measuring 160 A in width and 90 A in diameter. When stained with uranyl acetate and viewed face on, it is found to be stained homogeneously; when viewed edge on, it appears as a trilaminar structure with two dense outer lines enclosing a less dense center. Several manipulations indicate that the entire ribbon is trilaminar, and the infrequency of this appearance 1s due to the tendency of LPS to orient itself on its widest surface. Thus, after dispersion by sonication, detergent or ether, a,nd in sectioned specimens, trilaminar structures are much more prominent. The trilaminar appearance is reminiscent of that of cross-sections of arrays of bimolecular leaflets such as mitochondrial mesolager or lecithin micelles (Green & E’leisher, 1963), and all of the structures visualized in wild-strain LPS can be intrrpreted as representing arrays of bimolecular leaflets (Plate XIV). The flat disc result’ing from sonication or ether treatment (A) is a trilaminar structure viewed face on, and is a macromolecular complex derived from the ribbon (C). The occasional donutshaped particle is difficult to interpret, but may represent a trilaminar structure with faces of unequal diameter (B). ‘Thtbre is good evidence that the observed structure is not due to artifacts which might be introduced at one of several points ; first in the extraction itself; second, in the staining procedure; and third, in the drying of the material on a grid. Evidence a.gainst the first possibility is the similar structure demonstrated in LPS derived by two different techniques, the phenol-water and the aqueous ether procedures. It is also doubtful that the staining procedure introduces artifacts, since the appearance of LPS was the same whether or not the material was fixed with buffered osmic aci(1 prior to staining or whether it was stained in distilled water, O-1 N-NaOH, or OGY AI-Tris buffer. In addition, a lead citrate stain also revealed similar structure, although there was some tendency towards aggregation (Plate XV). An artifact due to severe drying is also unlikely, since embedded and sectioned LPS had an appearance similar to that, which was air-dried on the grid. Tho data indicate that the major determinant of the morphology of lipopolysaccharide is the lipid moiety. As mentioned previously, the appearance and dimensions are suggestive of phospholipid micelles or bimolecular leaflet-like structures. A similar appearance was found regardless of the polysaccharide content of the LPS. LPS from the heptoseless mutant, which is low in polysaccharide content, formed trilaminar structures similar to those found in wild-type LPS, which is 60% carbohydrate by weight. The dispersion of LPS by detergent and lipid solvent and tht demonstrated compression and ex-pansion of t’he trilaminar structure also suggest t,he importance of the lipid component in the morphology of LPS. Moreover, the behavior of LPS during sonication parallels that reported for lecithin aggregates when studied by light-scattering techniques. Attwood & Saunders (1965) found that sonication reduced asymmetrical lecithin aggregates to smaller, relat,ively stable, symmetrical, elongated spheroids with a, molecular weight of 2 :.: 10”. ItI our at.udy, sonication of LPS in aqueous suspt>nsion resulted in a rapid fall in
20
J. TV. SHANDS,
JR.,
J. A. GRAHAM
AND
Ii.
NATH
optical density and a relatively stable dispersion. Morphologically, sonication reduced the large aggregates to fairly uniform disc-like particles with an average diameter twice that of the unsonicated ribbon, namely, 320 A versus 160 A. A calculation of the molecular weight of these particles, estimating a density of 1.3 based on the chemical composition, gives a figure of approximately 6.0 x 105, which is in fair agreement with the above mentioned value for lecithin. If the most important determinant of the structure of lipopolysaccharide is the lipid moiety, where is the polysacoharide ? The data permit some interpretations but they are by no means conclusive. The observation that the trilaminar structures seen in LPS from the heptoseless mutant are smaller and have more delicate outer lines than that of both the wild strain and the RI1 mutant suggests that the “core” polysaccharide contributes to the trilaminar structure, perhaps to the dense lines. Since the RI1 mutant LPS has dimensions identical with that of the wild strain, it appears that the 0-polysaccharide side-chains do not contribute to the visible structure. Moreover, some evidence suggests that they are external to it. First, wild-strain LPS never packs together, whereas ribbons of LPS from the RI1 mutant may lie in close proximity (Plate XI). Second, in Plate VI the combination of positive and negative staining reveals a relatively clear area around most of the trilaminar structures, suggesting the presence of some material which does not stain; and third, the preparation of LPS labeled with ferritin-conjugated antibody shows ferritin lying up to 5008 from the ribbons (see Plate VIII). The biological significance of this trilaminar structure common to the preparations of lipopolysaccharide is obscure. It is possible that it may take some role in the diverse activities of Gram-negative LPS, and in this regard, it is of interest that the lipid moiety of LPS has attracted attention in the past as the possible toxigenic portion of the molecule (Westphal et al., 1958). The relative non-toxicity of LPS from a heptoseless mutant of Salmonella (Kessel, Freedman 6 Braun, 1966) renders this possibility unlikely, but further investigation is needed since this material may lack toxicity because of its lability. In our hands, the trilaminar structure of heptoseless LPS disappears even upon lyophilization. Moreover, we have not found a toxic preparation of LPS that lacked a trilaminar structure. The bimolecular leaflet-like structure of LPS suggests that it may be identical with the outer membrane of Gram-negative bacteria. Evidence that favors this possibility was found in electron microscopic studies of Veillonella parvulu before and after extraction with phenol (Bladen & Mergenhagen, 1964; Mergenhagen, Bladen & Hsu, 1966), and further support was provided by Rothfield et al. (1966) who observed the incorporation of LPS into phospholipid micelles. The latter authors proposed that LPS may exist in the outer membrane of a bacterium as a bimolecular leaflet interspersed or continuous with the membrane leaflets. The final solution to this problem concerning the relationship between LPS and Gram-negative cell wall will require much additional study; but regardless of the outcome, lipopolysaccharide may provide an interesting model for structural studies of biological membranes, particularly in the light of their remarkable stability. The technical assistance of Mrx L. Brown and Mr W. C. Herbert is gratefully acknowledged. We are indebted to Dr P. Small for his assistance in the preparation of the manuscript. The investigation \vas supporLed by U.S. Public Heitlth Service Grant AI 1302 10 from
the National
Institute
of Allergy
and Infectious
Disease.
LIPOPOLYSACCHARIDE
MORPHOLOGY
21
REFERENCES Attwood, D. & Saunders, L. (1965). Biochim. biophys. Acta, 98, 344. Beer, H., Braude, A. I. & Brinton, C. C., Jr. (1966). Ann. N.Y. Acud. Sci. 133, 450. Bladen, H. A. & Mergenhagen, S. E. (1964). J. Bach 88, 1482. Green, D. E. & Fleischer, S. (1963). Biochivn. bio@ys. Acta, 70, 654. Herzberg, M. & Green, J. H. (1964). J. Gen. Microbial. 35, 421. Kessel, R., Freedman, H. L Braun, W. (1966). J. Butt. 92, 592. Liideritz, O., Galanos, C., Risse, H. J., Ruschmann, E., Schlecht, S., Schmidt, G., SchulteHolthausen, H., Wheat, R., Westphal, 0. & Schlosshardt, J. (1966). Ann. N. Y. Acad. sci.
133, 349.
Mergenhagen, S. E., Bladen, H. A. & Hsu, K. C. (1966). Ann. N.Y. Acud. Sci. 133, 279. Mimer, K. C., Anacker, R. L., Fukushi, K., Haskins, W. T., Landy, M., Malmgren, B. & Ribi, E. (1963). Back Rev. 27, 352. Nikaido, H., Naide, Y. & M&k&, P. H. (1966). Ann. N.Y. Acad. Sci. 133, 299. Osborn, M. J. (1966). Ann. N.Y. Acud. Sci. 133, 375. Ribi, E., Mimer, K. C. & Perrine, T. D. (1959). J. Iwmunol. 82, 75. Rothfield, L., Takeshita, M., Pearhnan, M. 8r.Horne, R. (1966). Fed. Proc. 25, 1495. Schramm, G., Westphal, 0. & Liideritz, 0. (1952). 2. Nuturj. 7b, 594. Shands, J. W. (1965). J. Bat. 90, 266. Stocker, B. A. D., Wilkinson, R. & Mlikelii, P. (1966). Ann. 1V.Y. Acud. Sci. 133, 331. Weidol, JV., Frank, H. & Martin, H. H. (1960). J. Fen. Microbial. 22, 158. Westphal, O., Liideritz. 0. & Bister, F. (1962). 2. Naturf. 7b, 148. Westphal, O., Nowotny, A., Liideritz, O., Hurni, H., Eichenberger, E. & Schonholzer, G. (1958). Pharm. Acta Helv. 33, 401.
The Morphologic Structure of Isolated Bacterial Lipopolysaccharide J. W. Department
SHANDS,
J. A. GRAHAM
JR.,
of Microbiology,
University
Gainesville,
AND
K. NATH
of Florida College of ilfedicine
Florida,
U.S.A.
(Received 17 October, 1966) The morphology of lipopolysaccharides isolated from smooth and rough strains of Salmonella typhimurium was studied by electron microscopy using a positivestaining procedure. When viewed face on, lipopolysaccharide from the smooth strain was a homogeneously stained ribbon which could be broken down to homogeneously stained discs by sonication or agitation with ether. When viewed edge on, the ribbon and disc had a trilaminar appearance reminiscent of cross-sections of arrays of bimolecular leaflets. Lipopolysaccharide from a mutant lacking Opolysaccharide side-chains had a similar appearance and identical dimensions, whereas that from a heptoseless mutant, lacking 0 and “core” polysaccharide had smaller and more delicate trilaminar structures. The behavior of lipopolysaccharides in lipid solvent and the common morphological feature found in lipopolysaccharide regardless of the polysaccharide content suggest that the major morphological determinant is the lipid moiety. There is suggestive evidence that the “core” polysaccharide contributes to the trilaminar structures, whereas the 0-polysaccharide side-chains do not.
1. Introduction The lipopolysaccharides of Gram-negative bacteria are macromolecular complexes which possess both endotoxic and 0-antigenic activities. Chemically these substances are composed of a lipid moiety (lipid A) and a polysaccharide moiety, the proportions of which differ from one species to another and even among different strains of the same species. Recent investigations of LPSt f rom Salmonella (Nikaido, Naide $ MLkelS, 1966; Liideritz et al., 1966; Osborn, 1966)indicatethatthelipopolysaccharides are composed of repeating sugar units which impart the antigenic specificity. These are attached to a polysaccharide “core” possessing a “backbone” of heptose phosphate and 2-keto-3-deoxyoctonate, which in turn is linked to lipid A. Morphologically, in electron micrographs of shadowed specimens, LPS from Escherichia coli has the appearance of 100 A droplets at neutral pH or long ribbons at pH 10 (Schramm, Westphal & Liideritz, 1952). Similar morphology was demonstrated in LPS from a strain of E. coli (Weidel, Frank & Martin, 1960) and in that from Bordetella pertussis (Milner et al., 1963). In a detailed study of Boivin-type LPS from E. coli Beer, Braude & Brinton (1966) found a variety of shapes: snake-like, donut-like, flat sheets and slender rod-like filaments. The latter particles possessed the t Abbreviation
used: LPS. lipopolyaaccharidp. 15
IA
J. IV.
SHANDS,
all..
J. 11. (:I<.-\H;\N
ASD
I<. SAi’I’H
greatest toxicity, indicating t,hat, t,hey. might, bc responsible for the tosicif,y of ihe crude extract. Attempts to correlate the biochemical and morphological data ha.ve led t’o t’ht: proposal of two possible models. On one hand, Milner et rtl. (1963) suggested that LPS is a cellulose-like “micellar structure” composed of aggregated bundles of haptenic polysaccharide. On the other hand, Rothfield, Takeshita, Pearlman & Horne (1966) proposed that LPS forms a phospholipid-like, ordered “leaflet structure”. This paper is concerned with additional observations of the morphology of LPS in an attempt to correlate morphology and chemical structure. Advantage is taken of the afEnity of LPS for certain heavy-metal stains which delineate structure not seen in specimens prepared by the shadow-cast technique. In addition, the recent availability of strains of Gram-negative bacteria deficient in one or more parts of their lipopolysaccharide now allows one to correlate specific structural changes with chemical differences. Purified LPS from a wild strain of Salmonella typhimurium was compared with those of two mutant strains, one lacking the 0-polysaccharide side-chains, the other lacking both side-chains and “core” polysaccharide.
2. Materials and Methods (a) Bacteriu A smooth strain of Salmonella ty$Gmurium (strain 7) was obtained from Dr M. Herzberg (Dept. of Bacteriology, University of Florida). The surface characteristics and cell wall composition of this bacterium have been described (Herzberg & Green, 1964). The mutants of S. typhimurium LT2, kindly provided by Dr B. A. D. Stocker (Dept. of Microbiology, Stanford University), were (1) TV1 19, an RI1 mutant, which possesses a lipopolysaccharide devoid of specific “0” side-chains, and (2) SL1102, a variant, the LPS of which contains no heptose. The lipopolysaccharide of this latter strain is composed of lipid A and &-k&o3-deoxyoctonate, the latter being the only carbohydrate component detected (Stocker, Wilkinson & M&e%, 1966). E. co&i 0113 was obtained from Dr E. Ribi (Rocky Mountain Laboratories, Hamilton, Montana). The smooth strains of S. typhimurium and E. coli were cultivated in glucose-mineral salts medium for 18 hr at 37’C on a shaker while the mutants were grown in Brain Heart Infusion Broth (D&o) under the same conditions. The bacteria were harvested by cent*rifugation and washed twice with distilled water. (b) Preparation
of cell walk
After washing, the bacteria were suspended in a small volume of water and disrupted in a French pressure cell at 14,000 lb/sq. in. The broken and intact cells were separated from the protoplasm by centrifugation at 27,000 g for 15 min. The pellet was resuspended in O-1 M-phosphate buffer (pH 8-O), and 10 mg of powdered trypsin (Difco) was added for each ml. of pellet. Following incubation with gentle stirring for 10 hr at 37”C, the walls were washed repeatedly in 1.0 M-N8c1 and in distilled water. Walls were separated from intact bacteria by gently resuspending them and by discarding the dense button which formed at the bottom of the tube during centrifugation. Washing was continued until the cell walls were judged clean by electron microscopy.
(c) Extraction
of lipopolysaccharide
LPS was extracted from cell walls by the procedure of Westphal, L;ideritz dt Bister (1952). Approximately 600 mg of cell walls were suspended in 100 ml. of distilled water and an equal volume of liquid phenol was added. The mixture was stirred for 30 min at 60°C following which the phenol and water phases were separated by chilling and centrifugation, and the water phase was removed. The procedure was repeated after the addition of 100 ml. of water to the phenol phase. The two water phases were combined, dialyzed in dist)illed water and lyophilized.
PLATE III. x 330,000.
LPS
from
8. typhimuriwn
strain
i stwind
with
umnyl
acetate.
x 100.000:
inxct,
PLATE XIV. Diagram showing possible intwpretati?n of’ various image+ ohsrrverl in micrographs in terms of bimolecular leaflet structure. A. Flat disc (observed after sonic&ion or ether treatment) is a trilaminar structure vicwcd face on. 8. Occasional donut-shaped part,irle may rnprrsent trilaminar structure with ~~rt~y~al faces. (‘. Twisted ribbon strurt,urr.
LIPOPOLPSdCCHARIDR
~IORPHOLOGT
17
(d) Electron microscopy Lipopolysaccharide was suspended in glass-distilled water at a concentration of 1 mg/ml. LPS extracted from S. typhimurium 7 was stained positively by adding a drop of 2% in-any1 acetate to 0.3 ml. of the suspension. A drop of this mixture was placed on a carboncoated grid and then quickly removed by touching filter paper to the edge of the drop. Since this procedure precipitated LPS derived from the rough strains, these materials were stained by inversion of a grid containing dried LPS onto a drop of many1 acetate for 1 I,O 8 min, followed by gentle washing with water. Shadow-casting was carried out with uranium oxide in an Hitachi HUS-313 vacuum evaporator. Sectioned LPS from strain 7 was prepared by precipitation with ferritin-labeled antibody, the preparation and specificity of which have been described (Shands, 1965). The precipitate was washed 3 times in 0.1 M-phosphate buffer (pH 7.2) and fixed for 1 hr in 1 ‘J; osmic acid using the same buffer. The precipitate was then pelleted in 1% bovine serum albumin in a small polyethylene tube, the supernatant fluid was aspirated from the pellet, and a drop of 25% glutaraldehyde was added to coagulate the protein. The coagulum of LPS in albumin was cut into small blocks, dehydrated in alcohol, and embedded in Epon 812. Thin sections were cut with the Porter-Blum MT-2 microtome (Ivan Sorvall, Norwalk, Conn.) and stained with uranyl acetate. The materials were examined with a Siemens Ehniskop I electron microscope. ,Most pictures were taken at a plate magnification of 40,000. Dimensions were determined by means of a Leitz microscope equipped with a micrometer eye-piece.
3. Results Plate I shows positively-stainedlipopolysaccharide extracted from S. typhimurium 7. The appearance is that of a flat ribbon which is primarily linear but which branches freely. Its diameter varies but averages 160 A. At infrequent intervals the ribbon narrows to 90 A, and in such places (indicated by arrows) there is a triple-layered structure which consists of two dense outer lines enclosing a less dense cent’er. The general morphology depicted in Plate I is consistent with that demonstrated by others with shadow-casting. However, as additional evidence that the large structures stained are LPS and that others are not missed by the staining procedure, the same preparation was stained with uranyl acetate and then shadowed lightly with uranium oxide. The result is represented in Plate II, which shows that shadow-casting does not expose any macromolecular structures which are not also stained. The trilaminar structure revealed in Plate I raises at least two possibilities. Since its appearance is infrequent, it is possible that it is introduced into the LPS at different points. An alternate possibility is that the entire ribbon is trilaminar, but because of asymmetry tends to rest on its widest surface. The trilaminar appearance would, therefore, be visible only at twists in the ribbon or when the ribbon was made to stand on its thin edge. Several observations indicate that this latter possibility is the correct one. Plate III shows stained LPS and an enlargement of a twist. A distinct trilaminar structure travels from one edge of the flat ribbon to the opposite edge. Plates IV through VII show the effects of dispersing LPS. The material in Plate IV was dispersed by sonication in a Branson Sonifier at 20 kc/s at maximum power for 3 to 5 minutes. It is evident that the long ribbons have been broken up into smaller particles. Some of these are flat, homogeneously stained discs with diameters up to 500 A (A), whereas many other particles (B) have a trilaminar appearance. The latter probably represent flat discs standing on edge, since some particles seem to be resting at an angle and have both trilaminar and disc appearance (C).
IS
J. IV.
SHANDS,
JR..
J. A. C;RA IHAM
ASI)
K.
S.i’l‘I-I
Plate V shows LPS dispersed with 0.5% Tween 80 for 10 minutes at 20°C and stained in the usual manner. Two effects are noteworthy. First, t’he long ribbons arc broken into shorter segments; and second, t,he width of the ribbon is reduced from 160 to 116 8. The width and thickness, therefore, approximate one another: namely. 116 x x 94 .&. With these changes, one might expect, to see more trilnminarstructure, as iti evident in Plate V. =\fter the dispersion in Tween 80, LPS is often trapped in droplets of Tween. Plate VI illustrates this situation and demonstrates that the dimensions of the LPS trilaminar structures may vary. In some areas they appear compressed, and in others they appear expanded. Plate VII shows a preparation of LPS dispersed by shaking it vigorously for several minutes in water saturated with diethyl ether. The result is quite similar to that of sonic dispersion. One finds flat discs (A) and occasional donut-sha,ped particles (C). In thicker areas of the preparation, there are numerous trilaminar structures and occasional particles which seem to be resting at an angle which appear as trilaminar st*ructures and discs (B). Jf LPS is constructed entirely as a trilaminar structure, and if this appears infrequently because LPS tends to lie on its widest surface, then in sections of LPS, where the orientation is random, one should see much more trilaminar structure. Plate VIII shows a lipopolysaccharide preparation which has been precipitated with ferritin-conjugated antibody, embedded, sectioned and stained. The amount of trilaminar structure visible is obviously increased. The structures demonstrated so far are from a phenol-water extract of cell walls of S. typhimurium 7. To rule out the possibilities that the structures are artifacts proa duced by the extraction procedure or are found only in extracts of Xalmonellu, milder extraction procedure was used on whole cells of S. typhimurium 7 and E. coli 0113, namely, the aqueous ether method of Ribi, Milner & Perrine (1959). Plates IX and X show the LPS extracted from E. coli and X. typhimurium, respectively. Although neither of the preparations appears morphologically as well defined as the phenol ext)ract of cell walls, similarities are easily identified. Both of these preparations contain trilaminar structures and one contains the familiar ribbon. The amorphous appearance of the remainder of the materials may be due t,o contaminants or t,o the higher protein content of aqueous ether extracts which ma,y alter the morphology. Lipopolysaccharide extracted from rough mutants of S. typhimurium which lack the 0-polysaccharide side-chains and both side-chains and “core” polysaccharide were also found to contain trilaminar struct#ures. Plate XI shows a stained sample of LPS from the RI1 mutant TV119. Although the material tended to clump, many triplelayered structures are visible. In spite of the fact that this LPS possesses no Opolysaccharide side-chains (Stocker, et al. 1966), its dimensions are the same as those of t’he wild strain 7 LPS. Plate XII shows LPS derived from strain SL1102 which lacks both side-chains and “core” polysaccharide. Like the other preparations, it had been lyophilized and reconstituted in distilled water. Peculiarly, its morphology is unlike that of the other preparations, since it has an appearance which suggests folded and indented vesicles. This appears, however. to have been the result of lyophilization, since a sample of the same preparation which had not been lyophilized produced many triple-layered structures (Plate XIII). A comparison of these trilaminar structures with those in LPS of the wild strain and RI1 mutant shows that they
LIPOPOL~~ACCHAI~IUE
AIoL~L’HOI,C)GY
I9
arc more delicate, having less dense outer lines and a smaller over-all dimension, namely, 60 A versus 90 A.
4. Discussion The illustrations show that lipopolysaccharide derived from a smooth strain of &. typhimurium by the phenol-water procedure is a ribbon-like structure measuring 160 A in width and 90 A in diameter. When stained with uranyl acetate and viewed face on, it is found to be stained homogeneously; when viewed edge on, it appears as a trilaminar structure with two dense outer lines enclosing a less dense center. Several manipulations indicate that the entire ribbon is trilaminar, and the infrequency of this appearance 1s due to the tendency of LPS to orient itself on its widest surface. Thus, after dispersion by sonication, detergent or ether, a,nd in sectioned specimens, trilaminar structures are much more prominent. The trilaminar appearance is reminiscent of that of cross-sections of arrays of bimolecular leaflets such as mitochondrial mesolager or lecithin micelles (Green & E’leisher, 1963), and all of the structures visualized in wild-strain LPS can be intrrpreted as representing arrays of bimolecular leaflets (Plate XIV). The flat disc result’ing from sonication or ether treatment (A) is a trilaminar structure viewed face on, and is a macromolecular complex derived from the ribbon (C). The occasional donutshaped particle is difficult to interpret, but may represent a trilaminar structure with faces of unequal diameter (B). ‘Thtbre is good evidence that the observed structure is not due to artifacts which might be introduced at one of several points ; first in the extraction itself; second, in the staining procedure; and third, in the drying of the material on a grid. Evidence a.gainst the first possibility is the similar structure demonstrated in LPS derived by two different techniques, the phenol-water and the aqueous ether procedures. It is also doubtful that the staining procedure introduces artifacts, since the appearance of LPS was the same whether or not the material was fixed with buffered osmic aci(1 prior to staining or whether it was stained in distilled water, O-1 N-NaOH, or OGY AI-Tris buffer. In addition, a lead citrate stain also revealed similar structure, although there was some tendency towards aggregation (Plate XV). An artifact due to severe drying is also unlikely, since embedded and sectioned LPS had an appearance similar to that, which was air-dried on the grid. Tho data indicate that the major determinant of the morphology of lipopolysaccharide is the lipid moiety. As mentioned previously, the appearance and dimensions are suggestive of phospholipid micelles or bimolecular leaflet-like structures. A similar appearance was found regardless of the polysaccharide content of the LPS. LPS from the heptoseless mutant, which is low in polysaccharide content, formed trilaminar structures similar to those found in wild-type LPS, which is 60% carbohydrate by weight. The dispersion of LPS by detergent and lipid solvent and tht demonstrated compression and ex-pansion of t’he trilaminar structure also suggest t,he importance of the lipid component in the morphology of LPS. Moreover, the behavior of LPS during sonication parallels that reported for lecithin aggregates when studied by light-scattering techniques. Attwood & Saunders (1965) found that sonication reduced asymmetrical lecithin aggregates to smaller, relat,ively stable, symmetrical, elongated spheroids with a, molecular weight of 2 :.: 10”. ItI our at.udy, sonication of LPS in aqueous suspt>nsion resulted in a rapid fall in
20
J. TV. SHANDS,
JR.,
J. A. GRAHAM
AND
Ii.
NATH
optical density and a relatively stable dispersion. Morphologically, sonication reduced the large aggregates to fairly uniform disc-like particles with an average diameter twice that of the unsonicated ribbon, namely, 320 A versus 160 A. A calculation of the molecular weight of these particles, estimating a density of 1.3 based on the chemical composition, gives a figure of approximately 6.0 x 105, which is in fair agreement with the above mentioned value for lecithin. If the most important determinant of the structure of lipopolysaccharide is the lipid moiety, where is the polysacoharide ? The data permit some interpretations but they are by no means conclusive. The observation that the trilaminar structures seen in LPS from the heptoseless mutant are smaller and have more delicate outer lines than that of both the wild strain and the RI1 mutant suggests that the “core” polysaccharide contributes to the trilaminar structure, perhaps to the dense lines. Since the RI1 mutant LPS has dimensions identical with that of the wild strain, it appears that the 0-polysaccharide side-chains do not contribute to the visible structure. Moreover, some evidence suggests that they are external to it. First, wild-strain LPS never packs together, whereas ribbons of LPS from the RI1 mutant may lie in close proximity (Plate XI). Second, in Plate VI the combination of positive and negative staining reveals a relatively clear area around most of the trilaminar structures, suggesting the presence of some material which does not stain; and third, the preparation of LPS labeled with ferritin-conjugated antibody shows ferritin lying up to 5008 from the ribbons (see Plate VIII). The biological significance of this trilaminar structure common to the preparations of lipopolysaccharide is obscure. It is possible that it may take some role in the diverse activities of Gram-negative LPS, and in this regard, it is of interest that the lipid moiety of LPS has attracted attention in the past as the possible toxigenic portion of the molecule (Westphal et al., 1958). The relative non-toxicity of LPS from a heptoseless mutant of Salmonella (Kessel, Freedman 6 Braun, 1966) renders this possibility unlikely, but further investigation is needed since this material may lack toxicity because of its lability. In our hands, the trilaminar structure of heptoseless LPS disappears even upon lyophilization. Moreover, we have not found a toxic preparation of LPS that lacked a trilaminar structure. The bimolecular leaflet-like structure of LPS suggests that it may be identical with the outer membrane of Gram-negative bacteria. Evidence that favors this possibility was found in electron microscopic studies of Veillonella parvulu before and after extraction with phenol (Bladen & Mergenhagen, 1964; Mergenhagen, Bladen & Hsu, 1966), and further support was provided by Rothfield et al. (1966) who observed the incorporation of LPS into phospholipid micelles. The latter authors proposed that LPS may exist in the outer membrane of a bacterium as a bimolecular leaflet interspersed or continuous with the membrane leaflets. The final solution to this problem concerning the relationship between LPS and Gram-negative cell wall will require much additional study; but regardless of the outcome, lipopolysaccharide may provide an interesting model for structural studies of biological membranes, particularly in the light of their remarkable stability. The technical assistance of Mrx L. Brown and Mr W. C. Herbert is gratefully acknowledged. We are indebted to Dr P. Small for his assistance in the preparation of the manuscript. The investigation \vas supporLed by U.S. Public Heitlth Service Grant AI 1302 10 from
the National
Institute
of Allergy
and Infectious
Disease.
LIPOPOLYSACCHARIDE
MORPHOLOGY
21
REFERENCES Attwood, D. & Saunders, L. (1965). Biochim. biophys. Acta, 98, 344. Beer, H., Braude, A. I. & Brinton, C. C., Jr. (1966). Ann. N.Y. Acud. Sci. 133, 450. Bladen, H. A. & Mergenhagen, S. E. (1964). J. Bach 88, 1482. Green, D. E. & Fleischer, S. (1963). Biochivn. bio@ys. Acta, 70, 654. Herzberg, M. & Green, J. H. (1964). J. Gen. Microbial. 35, 421. Kessel, R., Freedman, H. L Braun, W. (1966). J. Butt. 92, 592. Liideritz, O., Galanos, C., Risse, H. J., Ruschmann, E., Schlecht, S., Schmidt, G., SchulteHolthausen, H., Wheat, R., Westphal, 0. & Schlosshardt, J. (1966). Ann. N. Y. Acad. sci.
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Mergenhagen, S. E., Bladen, H. A. & Hsu, K. C. (1966). Ann. N.Y. Acud. Sci. 133, 279. Mimer, K. C., Anacker, R. L., Fukushi, K., Haskins, W. T., Landy, M., Malmgren, B. & Ribi, E. (1963). Back Rev. 27, 352. Nikaido, H., Naide, Y. & M&k&, P. H. (1966). Ann. N.Y. Acad. Sci. 133, 299. Osborn, M. J. (1966). Ann. N.Y. Acud. Sci. 133, 375. Ribi, E., Mimer, K. C. & Perrine, T. D. (1959). J. Iwmunol. 82, 75. Rothfield, L., Takeshita, M., Pearhnan, M. 8r.Horne, R. (1966). Fed. Proc. 25, 1495. Schramm, G., Westphal, 0. & Liideritz, 0. (1952). 2. Nuturj. 7b, 594. Shands, J. W. (1965). J. Bat. 90, 266. Stocker, B. A. D., Wilkinson, R. & Mlikelii, P. (1966). Ann. 1V.Y. Acud. Sci. 133, 331. Weidol, JV., Frank, H. & Martin, H. H. (1960). J. Fen. Microbial. 22, 158. Westphal, O., Liideritz. 0. & Bister, F. (1962). 2. Naturf. 7b, 148. Westphal, O., Nowotny, A., Liideritz, O., Hurni, H., Eichenberger, E. & Schonholzer, G. (1958). Pharm. Acta Helv. 33, 401.