Ultracentrifugation as a Means for the Separation and Identification of

Chapter 12

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Ultracentrifugation as a Means for the Separation and Identification of Lipopolysaccharides Marshall Phillips and Kim A. Brogden Agricultural Research Service, U.S. Department of Agriculture, National Animal Disease Center, P.O. Box 70, Ames, IA 50010 Ultracentrifugation can be used as a method to separate and identify different types of lipopolysaccharides (LPS). To illustrate this, LPS from Pasteurella multocida. Brucella abortus, and other gram-negative microorganisms were fractionated on 34% to 45% CsCl field-formed gradients. After centrifugation, their apparent buoyant densities ranged from 1.34 to 1.44 gm/ml. The buoyant density positions in the gradients are thought to be not only a function of their chemical composition, but also of the hydration and conformation of the LPS molecules in their respective solutions. These differences can be capitalized on for the isolation of contaminant-free LPS as well as the isolation of LPS with slightly differing molecular composition. This may prove to be an ideal way to separate and recover recombinant LPS for biotechnology purposes. The LPS can then be used directly for the development of immunizing and diagnostic reagents.

Lipopolysaccharide (LPS) is an integral component of endotoxin found at the outer surface of gram-negative bacteria. This moiety is responsible for much of the toxicity ascribed to endotoxin and is an important determinant involved in the pathogenesis of gram-negative bacterial infections. Chemically, monomeric molecules of LPS have a molecular weight of about 2 kilodaltons, and are comprised of a hydrophilic region of polysaccharide (somatic polysaccharide of repeating units up to 100 monosaccharides long) and a hydrophobic region of long chain (n-14, n-20) fatty acids linked to N-acetylglucosamine (Figure 1). The two regions are joined by a "core" polysaccharide containing 2-keto-3-deoxyoctulosonic acid (KDO), heptose, and amino sugars. Ethanolamine and phosphate groups are covalently attached. Mutant bacterial strains exist that lack components of the core polysaccharide (Ra through Re). Excellent reviews have been written on the composition of LPS (1, 2) including its components such as lipid A (3) and KDO (4). In this paper, we present ultracentrifugation both as a preparative technique to separate and purify different types of LPS and This chapter not subject to U.S. copyright Published 1990 American Chemical Society In Downstream Processing and Bioseparation; Hamel, J., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1990.

12. PHILLIPS & BROGDEN

Ultracentrifugation and Lipopolysaccharides

239

GALACTOSE


RHAMNOSE φ

SOMATIC POLYSACCHARIDE

Ra

MANN0SE + ABEQUOSE

CORE POLYSACCHARIDE

LIPID A

CORE POLYSACCHARIDE

LIPID A

Rb

6AL-GLU-HEP-KD0

Rc

, GLU-HEP-KDO

Rd

|

LIPID A

LIPID A

H E P - Κ DO

LIPID A

Re , KDO

LIPID A

Figure 1. A schematic diagram illustrating a basic lipopolysaccharide molecule from Salmonella t y p h i m u r i u m with the polysaccharide and lipid regions. Also shown are core polysaccharide mutants (Ra through Re).

In Downstream Processing and Bioseparation; Hamel, J., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1990.


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as an analytical technique to assess the purity of LPS preparations and to distinguish L P S of slightly differing chemical composition. To illustrate this, L P S from different gram-negative bacteria as w e l l as L P S f r o m Salmonella minnesota Ra, Rc, and Re mutants were fractionated on 34% to 45% C s C l gradients. Differences i n density of these LPS can be capitalized on for their isolation, purification, and characterization. Equilibrium (isopycnic) centrifugation is well established (5-8). The basic concept involves the generation of a gradient during centrifugal force: the density near the bottom of the centrifuge tube is greater than that of the most dense particle i n the mixture and, at the meniscus, the density is less than that of the lightest particle. The initial application of the technique was applied to the characterization of nucleic acids, and is of historical importance i n the discovery of D N A replication (9). Proteins and glycoproteins have also been characterized w i t h this technique (10). However, there is a paucity of reports on the use of the technique for the study of LPS. Morrison and Leive (11) first used the technique to separate LPSs w i t h s l i g h t l y d i f f e r i n g chemical composition extracted f r o m Escherichia coli. They observed that the chemical composition of the LPS was related to its apparent buoyant density and was due to the length of the somatic polysaccharide. A nearly direct correlation was observed between the amount of carbohydrate i n the LPS preparation and its buoyant density. Later, the biological properties of the LPS was also shown to correlate with the buoyant density (12-14). In 1978, Phillips and Rebers (15) published the first analytical equilibrium centrifugation patterns of LPS i n C s C l gradients. This report and subsequent reports (16-18) indicated, from the schlieren patterns, that well-defined molecular distributions of LPS molecules exist at equilibrium in C s C l gradients. A number of media may be employed for the preparation of gradients (5, 8). These include simple sugars and analogous polyhydroxyl compounds, polysaccharides, proteins, iodinated organic compounds (i.e. metrizamide), colloidal silica (Ludox), and inorganic salts (8). Inorganic salts, such as C s C l , are the most commonly used media for the preparation of gradients (8). The advantages of C s C l are numerous. First, C s C l is well characterized and its thermodynamic properties known. Second, C s C l is very soluble i n water resulting i n very dense solutions with a relative viscosity close to unity. T h i r d , pure C s C l solutions do not absorb ultraviolet light. Finally, high concentrations of C s C l (34% to 50%) are very chaotropic and provide conditions for the disruption of hydrogen bonding. Thus, under the conditions of C s C l equilibrium centrifugation, the apparent buoyant density positions of L P S w i l l be an e q u i l i b r i u m balance among the forces of hydrophobic interactions, solvation, and chaotropic-mediated actions. H o w e v e r , C s C l diffuses r a p i d l y and a preformed gradient is stable for only a short time.



Ultracentrifiigation and Lipopolysaccharides

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METHODS FOR EXTRACTION OF LPS Since L P S varies i n chemical composition from one bacterial species to another, different methods are used for its extraction. L P S is generally extracted from smooth strains of bacteria i n a mixture of phenol and water (PW) at 68°C as described by Westphal and Jann (19). After extraction, the solution is allowed to cool and partition. Nearly all proteins remain i n the phenol phase or interphase, while polysaccharides, LPS, and nucleic acids remain i n the aqueous phase. In some instances, the L P S from certain organisms (i.e. B r u c e l l a a b o r t u s ) r e m a i n i n the p h e n o l phase. Lipopolysaccharide is extracted from rough strains of bacteria i n a mixture of phenol, chloroform, and petroleum ether (PCP) at room temperature as described b y Galanos, et aL (20). After extraction, the chloroform and petroleum ether are removed by rotary evaporation and the L P S is then precipitated from the phenol with water. W i t h some bacteria, P W or P C P w i l l not extract sufficient quantities of LPS (21) and E D T A (22), chloroformmethanol (23), or butanol (11) must be used.

PREPARATIVE CsCL GRADIENT ISOLATION OF LPS After extraction, L P S solutions usually contain protein and nucleic acid contamination. These can be removed by a variety of procedures (24). The most commonly used procedures treat L P S solutions with nucleases and proteolytic enzymes. Proteinase Κ or proteinase Κ solutions containing sodium dodecyl sulfate (SDS) can also be used. Alternately, contaminantfree L P S can be obtained by preparative equilibrium centrifugation. In our work, crude preparations of LPS were suspended i n 0.05 M Tris buffer, p H 7.8, and added to solutions of C s C l i n the same buffer. The concentration of C s C l was adjusted such that the final concentration was 34% after adding 1 m l of LPS i n Tris buffer. The gradient was attained i n 38 hours at 39,000 χ g in the Beckman SW 50.1 rotor. Recovery of L P S from the gradient is accomplished b y sequentially emptying the tube for recovery of the L P S band with a Pasteur pipette. The C s C l is then removed by dialysis. In this technique, LPS buoy i n a range from 1.34 to 1.45 g m / c m i n C s C l gradients. Centrifugation was performed at 25°C to limit diffusion of the C s C l i n the tubes when manipulated at room temperature. Contaminants w i l l buoy at different densities. For example, the lipids w i l l buoy on the surface of the gradient, most proteins w i l l buoy above 1.34 g m / c m (usually 1.25 to 1.32 g m / c m ) , extraneous complex carbohydrates w i l l buoy from 1.49 to 1.58 g m / c m or higher, and nucleic acids w i l l band at even higher values (Figure 2). The buoyant density of these molecules i n C s C l is not only a function of their chemical content but also of their degree of hydration, conformation, orientation, and extent of hydrogen bonding. Procedures and calculations for obtaining equilibrium for other density regions are also described (6-8). 3

3

3

3



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Lipids 1.25

} Proteins

1.35 1.45 1.55 1.65 Figure 2. gradients.

J Lipopolysaccharides ^Carbohydrates

Nucleic Acids

Predicted buoyant position of hydrated constituents


in CsCl



243

The amount of carbohydrate i n the L P S molecule influences its buoyant density position. This can be shown by mixing Ra, Rc, and Re LPS forms from S a l m o n e l l a m i n n e s o t a and seperating them i n a 39% C s C l gradient (Figure 3). The respective forms buoy at 1.38, 1.42, a n d 1.50 g m / c m . Each LPS i n the Ra, Rc, and Re mixture separated at each density as clearly as each individual form alone. Similarly, representative patterns of Brucella L P S are shown i n Figure 4. The LPS reaches equilibrium i n a narrow region at one position, and other contaminants reach equilibrium at other positions i n the tubes. M i n o r contaminants m a y not be identifiable as a band but only detected i n isolated fractions by chemical means (i.e. protein determination, ultraviolet absorption, etc.). It is also possible that L P S molecules may bind with components and thus be found in other regions i n the gradient. Preparations of L P S isolated from Brucella abortus strains S-19 and 2308S band at nearly identical positions. The LPS from B . meletensis can be found i n two bands. The B. abortus rough strains contain a band at 1.35 g m / c m a n d , under conditions of the gradient, also can be found i n more than one band. Table I contains a listing of the apparent buoyant densities of L P S isolated from several sources.


3

3

ANALYTICAL CsCL GRADIENT SEPERATION OF LPS Analytical ultracentrifugation of LPS on C s C l gradients reveals a number of properties about the L P S molecule that cannot be easily determined by typical chemical analysis (i.e. protein, total carbohydrate, K D O content, etc.). A n a l y t i c a l ultracentrifugation is performed i n the Beckman M o d e l Ε analytical ultracentrifuge. D u r i n g centrifugation, a schlieren peak is generated. The shape and number of peaks present determine not only the purity of the L P S preparation but the number of L P S molecules w i t h slightly differing chemical composition. Therefore very sharp, narrow schlieren peaks indicate a monodisperse, h i g h l y u n i f o r m L P S , free of contamination whereas a wide, less defined peak indicates the presence of a population of differing LPS molecules of questionable purity. Also, the use of the analytical centrifuge can be used to characterize the L P S obtained by the preparative procedure. Methods for determining the apparent density values from the schlieren peaks are based o n the general equation of e q u i l i b r i u m centrifugation (9, 25-27). For example, a monodisperse schlieren peak was observed when B. abortus L P S was recovered from the discrete bands as shown i n preparative tube A , Figure 5. Monodisperse peaks are not always observed f r o m the B r u c e l l a L P S (Figure 6). Monodisperse peaks were also obtained w i t h L P S f r o m P a s t e u r e l l a multocida (Figure 6c). Figure 7 shows the computer-assisted conversion of components expected from schlieren patterns as described by Johnson (28).



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A

B

C


D

Figure 3. Buoyant density separation of Salmonella minnesota Ra, Rc, and Re lipopolysaccharides. Centrifugation is 39% C s C l for 40 hours at 25°C A : Ra; B: Rc; C : Re; and D : Mixture of Ra, Rc, and Re.

Figure 4. Preparative 36% and 39% C s C l gradients of Brucella LPS. Shows separation of other components. Left: 39% gradient showing B. meletensis LPS. Right: 36% gradient showing B. abortus LPS.


12i PHILLIPS & BROGDEN


245

Table I. Apparent Buoyant Densities of LPS

Origin

Apparent Density (g/cm )

Reference


3

Escherichia coli 0811 (phenol extraction) Escherichia coli 08111 (butanol extraction) Salmonella minnesota R595 Pasteurella multocida X-73 Pasteurella multocida Serratia marcescens Brucella abortus 2308 Brucella abortus S-19 Brucella abortus 2308R Brucella meletensis

1.45

11

1.35

11

1.38 1.39 1.38-1.44 1.50 1.35 1.35 1.35-1.38 1.38

12 16 16 13 17 17 -

Figure 5. 34% C s C l gradients of Brucella LPS. The LPS from B. abortus strain indicated under tube.



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AND BIOSEPARATION

Figure 6. M o d e l Ε analytical schlieren patterns of p u r i f i e d L P S . Equilibrium attained at 36,000 r p m at 25°C for 18 hours i n 2°C sector cells. A : 34% C s C l pattern of B. abortus L P S , sample from preparation r u n i n Figure 5 A . B: 34% C s C l pattern of B. abortus LPS. C: 39% C s C l pattern of P. multocida LPS.

Figure 7. Computer-generated density gradient distributions. N o t e first r o w : concentration of components; second r o w : schlieren pattern.


12. PHILLIPS & B R O G D E N

Ultracentrifugation and Lipopolysaccharides 247


EFFECT OF CHEMICAL TREATMENT OF LPS ON SEPARATION The buoyant density of LPS has been shown to differ between that extracted from smooth and rough bacteria (for example, B. abortus strain 2308, Figure 5), as well as differ within a single bacterial species depending u p o n its method of extraction (11), ionic environment, and association w i t h other molecules. The effect of chemical treatment on L P S was shown b y Amano and Fukushi (29). Treatment of S. minnesota w i l d and R mutants (Ra to Re) w i t h alkali caused partial removal of somatic polysaccharide linked fatty acids. Thus, the monolayer formation i n alkali-treated L P S resulted from reduction of the steric hindrance b y fatty acids and added to the reduction of its hydrophobicity. A t acidic p H , changes i n aggregation of L P S have also been observed. Following treatment with acetic acid ( p H 2.8) for 3 hours at room temperature, Acker and Wartenberg (30) observed L P S strands w i t h regularly spaced, "loosened" regions. After treatment for several days, the LPS strains were disconfigured. Divalent cations are important to the physical-chemical properties of LPS (31). The L P S contains inorganic cations such as N a K , C a , and M g , as well as l o w molecular weight amines (1). These cations neutralize the negative charges on LPS molecules necessary for ordered assembly (32). Treatment of LPS with detergents result i n a reversible dissociation of L P S into subunits (33). Both Ribi, et aL (34) and Horisberger and Dentam (35) observed that L P S treated w i t h sodium deocycholate, followed by dialysis, d i d not alter the size of the particles, yet resulted i n a more u n i f o r m preparation. P o l y m y x i n Β w i l l dissociate and degrade L P S components. These effects have been suggested to be caused b y the interactions of the lipophilic and lipophobic groups of the antibiotic with those of the LPS (36). A l l of these treatments m a y influence the density of L P S preparations b y removing contamination components that interact w i t h the L P S molecules. +

+

+ +

+ +

EFFECTIVENESS OF CsCL GRADIENT SEPARATION The preparative C s C l gradients effectively separate L P S from contaminants and the analytical C s C l gradients effectively characterize the L P S i n them. The hypersharp peak patterns indicate large aggregates (or highly associated molecules) of L P S under the conditions of the C s C l . This is the result of the association of the hydrophobic portions of the molecules into the formation of a highly oriented conformation. Based o n the expected patterns of molecules i n C s C l (Figure 6), the schlieren patterns indicate only a limited population distribution of molecules i n the preparations. In the gradients, it is thought that the L P S form a continuum of different size micelles. Since L P S are amphipathic molecules, they undergo association dissociation phenomena, and enter into micellular formation. The L P S molecules combine by hydrophobic - h y d r o p h i l i c interactions; l i p i d A

American Chemical Society Library 1155 16ih St., N.W. In Downstream Processing and Bioseparation; Hamel, J., et al.; Washington, DSL Society: 2 0 0 » Washington, DC, 1990. ACS Symposium Series; American Chemical

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DOWNSTREAM PROCESSING AND BIOSEPARATION

regions align toward each other internally, and their carbohydrate regions orientate toward the aqueous interphase. The nature of such aggregate formation is based on pH, temperature, and other solution conditions.


FUTURE APPLICATION Ultracentrifugation can be used as a method to separate and identify different types of lipopolysaccharides (LPS). The buoyant density positions of LPS in the gradients not only reveal something about their state of purity but also about their chemical composition. These differences can be capitalized on for the isolation of contaminant-free LPS as well as the isolation of LPS with slightly differing molecular composition. This may prove to be an ideal way to separate and recover recombinant LPS for biotechnology purposes. The LPS can then be used directly for a variety of biomedical purposes including immunomodulators, vaccines, and diagnostic reagents.

LITERATURE CITED: 1.

2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.

Luderitz, Ο.; Freudenberg, Μ. Α.; Galanos, C.; Lehmann, V.; Rietschez, E. T.; Shaw, D.E.; Current Topics in Membranes andTransport;Membrane Lipids of Prokaryotes: Razin, S.; Rottem, S., Eds.; Academic: New York, 1982; Vol. 17, pp. 79-151. Westphal, O.; Jann, K.; Himmelspach, K.; Prog. Allergy 1983, 33, pp. 9-39. Rietschel, E. T.; Brade, H.; Brade, L.; Brandenburg, K.; Schade, U.; Seydel V.; Zahringer, U.; Galanos, C.; Luderitz, O.; Westphal, O.; Labischinski, H.; Kusumoto, S.; Shiba, T.; Progr. Clin. Biol. Res. 1987, 231,pp.25-53. Unger, F. M.; Advances in Carbohydrate Chemistry andBiochemistry;Tipson, R. S.; Horton, D., Eds.; Academic: New York, 1981; pp. 323-88. Osterman, L. Α.; Equilibrium (Isopycnic) Centrifugation; Springer-Verlag: New York, 1984; pp. 275-307. Ifft, J. B.; A Laboratory Manual of Analytical Methods of Protein Chemistry: Alexander, P.; Lundgren, H. P.; Eds.; Pergamon: Oxford, 1969; Vol. 5, pp. 151. Hearst, J. E.; Schmid, C. W.; Methods in Enzymology: Hirs, C. H.; Timsahoff, S. N., Eds.; Academic: New York, 1973; Vol. 27, pp. 111-27. Birnie, G. D.; Rickwood, D.; Centrifugal Separations in Molecular and Cell Biology; Butterworths: London, Boston, 1978; pp. 169-217. Meselson, M.; Stahl, F. W.; Vinograd, J. ; Proc. Nat. Acad. Sci. U.S.A. 1957, 43, pp. 581-588. Dunstone, J. R.; Prep. Sci. 1969, 4, pp. 267-85. Morrison, D. C.; Leive, L.; T. Biol. Chem. 1975, 250, pp. 2911-19. Ulevitch, R. J.; Johnston, A. R.; Clin. Invest. 1978, 62, 1313-24. Wilson, M. R.; Morrison, D. C.; Eur.J.Biochem. 1982, 128, pp. 137-41. Vukajlovich, S. W.; Morrison, D. C. ;J.Immunol. 1985, 135, pp. 2546-50. Phillips, M.; Rebers, P. A. ; Anal. Biochem. 1978, 85, pp. 265-70. Rimler, R. B.; Rebers, P. Α.; Phillips, M.; Am. J. Vet. Res. 1984, 45, pp. 759-63. Phillips, M.; Pugh Jr, G. W.; Deyoe, B. L.; Am.J.Vet. Res. 1989, 50, pp. 311-317. Phillips, M.; Brogden, Κ. Α.; Infect. Immun. 1987, 55, pp. 2047-51.


12. PHILLIPS & BROGDEN 19. 20. 21. 22. 23. 24. 25. 26. Downloaded by UNIV OF MINNESOTA on October 2, 2013 | http://pubs.acs.org Publication Date: January 24, 1990 | doi: 10.1021/bk-1990-0419.ch012

27. 28. 29. 30. 31. 32. 33. 34. 35. 36.


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Westphal, O.; Jann, K.; Methods in Carbohydrate Chemistry; Whistler, R. L., Ed.; Academic: New York, 1965; pp. 83-92. Galanos, C.; Luderitz, Ο.; Westphal, Ο.; Eur. J. Biochem. 1969, 9, pp. 245-49. Wu, L.; Tsai, C-M.; Fresch, D. E.; Anal. Biochem. 1987, 160, pp. 281-89. Leive, L.; Methods in Enzymology; Ginsburg, V., Ed.; Academic Press: New York, 1972; Vol. 28, pp. 254-62. Amano, K.; Fukushi, F. ; Microbiol. Immunol. 1984, 28, pp. 135-148. Rietschel, E. T.; Handbook of Endotoxin; Chemistry of Endotoxin; Elsevier Science: The Netherlands, 1984; Vol. 1. Svedberg, T.; Pederson, K. O.; The Ultracentrifuge; Oxford Press: Oxford, 1940. Vinograd, J.; Hearst, J. E.; Prog. Chem. Organic. Natural Products 1962, 20, pp. 371-405. Chervenka, C. H.; A Manual of Methods for the Analytical Ultracentrifuge; Spinco Div. Beckman Instruments, California, 1969. Johnson, N. W.; J. Gen. Virol. 1973, 18, pp. 207-09. Amano, K.; Fukushi, F.; Microbiol. Immunol. 1984, 28, pp. 161-168. Acker, G.; Wartenberg, K.; Zbl.Bakt.Hyg. I Abt. Orig. A 1976, 235, pp. 439-52. Galanos, C.; Luderitz, Ο.; Eur.J.Biochem. 1975, 54, ΡΡ- 603-10. Graham, G. S.; Treick, R. W.; Brunner, D. P.; Curr. Microbiol. 1979, 2, pp. 339-43. Huribert, R. E.; Hurlbert, I. M.; Infect. Immun. 1977, 16, pp. 983-94. Ribi, E.; Anacker, R. L.; Brown, R.; Haskins, W. T.; Malmgren, B.; Milnder, K. C.; Rudbach, J. A. ;J.Bacteriol. 1966, 92, pp. 1493-1509. Horisberger, M.; Dentam, E.; Arch. Microbiol. 1980, 128, pp. 12-18. Weber, D. Α.; Nadakavukaren, J.; Tsang, J. C.; Microbios 1976, 17, pp. 149-61.

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Ultracentrifugation as a Means for the Separation and Identification of

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