inclusion bodies of prokaryotes

Copyright 1974. All rights reserved

INCLUSION BODIESOF PROKARYOTES

~1635

J. M. ShivelyDepartment of Biochemistry, Clemson University, Clemson, South Carolina 29631

CONTENTSINTRODUCTION ................................................................ 167NONMEMBRANE-ENCLOSED INCLUSIONS ...................................... 168

Polyglucoside (a, Glycogen) Granules ...................................... 168Polyphosphate (Volutin, Metachromatie) Granules ............................ 168Cyanophycin (Structured) Granules ........................................ 170Ph ycobilisomes ......................................................... 171Crystals and Paracrystalline Arrays ........................................ 172Tubules, Microtubules, and Raphidosomes .................................. 176

MEMBRANE-ENCLOSED INCLUSIONS ........................................... 176Polyglucoside (Glycogen) Granules ........................................ 176Poly-~-hydroxybutyrate Granules .......................................... ¯ 178Sulfur Globules ........................................................ 179Gas Vacuoles .......................................................... 179Polyhedral Bodies (Carboxysomes) ........................................ 180Chlorobium Vesicles .................................................... 182

OTHER INCLUSIONS ........................................................... 184CONCLUDING REMARKS ...................................................... 185

INTRODUCTION

This review summarizes our current knowledge of prokaryotic inclusion bodies. Theinclusions have been divided into two major groups based on the presence or absenceof a surrounding membrane. A third group (see section: OTHER INCLUSIONS)

includes those which, due to a lack of information, could not be classified.Comparisons of the inclusion bodies of prokaryotes and eukaryotes, evaluations

of experimental methodology, and a discussion of endospores (36, 42, 43, 54, 130)and ribosomes (70, 83) are not included. Only the structure of the inclusions re-vealed by electron microscopy is described, i.e. light microscope appearances are notpresented.

The literature cited has been reduced to what this author feels is a listing of themost pertinent references. When a recent review is available it is listed following the

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168 SHIVELY ~

section title, e.g. Polyphosphate Granules (46), and the reader is directed to thisreview for more details, background information, and literature citations.

NONMEMBRANE-ENCLOSED INCLUSIONS

Polyglucoside (a, Glycogen) Granules

Many prokaryotic organisms have been shown to store polymers of glucose (9, 14,24, 72, 86, 96, 139). The polyglucoside may be dispersed throughout the cytoplasmor deposited as membrane-enclosed (see below) or nonmembrane-enclosed granules(138).

The granules consist of highly branched, high molecular weight polymers whichresemble either glycogen or amylopectin (9, 24, 44, 72, 86, 96, 139). In the bacteria(Figure 1), the granules are 20-100 nm in diameter and commonly have an unevenappearance (14, 86). The granules of the blue-green algae may be crystals, spheres(Figur.e 2), or rods and are generally observed between the thylakoid membranes(139). The rods of Nostoc muscorum are 30 nm wide, 65 nm long, and consist oftwo equal parts (9); those of Oscillatoria rubescens (Figure 3) may be as long as 300nm and are composed of 7.0 nm discs with a central pore (62). In the blue-greenalgae the spherical granules are generally 25-30 nm in diameter (139). The maxi-mum size of the granule, at least those with a regular shape, may be governed bypacking limitations (92). The granules are commonly very electron dense in thinsections stained with lead citrate.

Bacteria produce the polymer in response to excess carbon when nitrogen, sulfur,or phosphorous are limiting, or when the pH is low (24,. 96). Granule formation the blue-green algae is correlated with active photosynthesis (72, 139). The polymeris hypothesized to be a storage form of energy and/or carbon. Synthesis in plantsand bacteria occurs by the following reactions (96):’

ATP q- ct-glucose-l-P ~ ADP-glucose -~- I ~ PiADP-glueose -I- a-l,4-gluean ~- ADP -t- a-1,4-glueosyl-glucan.

Polyphosphate (Volutin, Metachromatic) Granules (46)

Polyphosphates are widely distributed in prokaryotes (23, 31, 46, 57, 58, 72, 82, 110,124, 129, 138, 139). The bulk of the polyphosphate is present as linear, high molecu-lar weight molecules, e.g. greater than 500 residues per molecule have been reported(31, 46). Small linear molecules, as well as cyclic tri-, tetra-, penta-, and hexameta-

Figure 1 Polyglucoside granules (arrows) in Escherichia coil Except where noted marker barin all figures equals 100 nm. All micrographs are thin sections except where indicated. Cour-tesy of J. W. Greenawalt, Johns Hopkins University Medical School, Baltimore, Md.

Figure 2 Polyglucoside granules (PG), cyanophycin granules (CG), and gas vacuoles of Aphanizomenon flos-aquae. Courtesy of R. B. Wildman (137); with permission.

Figure 3 Isolated polyglucoside granules of Oscillatoria rubescens: negative stain. Courtesyof M. Jost, AEC-Michigan State University, Plant Research Laboratory, East Lansing.

Figure 4 Polyphosphate granules (arrows) in Thiobacillus novellus. J. M. Shively (113); withpermission.


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170 SHIVELY

phosphates have been found in some cases (46, 82). The polyphosphate may consti-tute 40-50% of the total cell phosphorous (31, 72).

The polymer is commonly deposited as spherical, electron-opaque granules (Fig-ure 4) which range in diameter from 48 nm to greater than 1 /~m; size depends onthe organism examined and on the state of granule development (31, 46, 56-58, 72,114, 139). The granules are usually located in the nucleoplasmic region of the cell(56, 57, 72, 139), but have been seen in association with other cell components (46,129). In many instances the granules reportedly evaporate in an electron-beam ofhigh intensity and assume a "holey" or mottled appearance (46, 139). Althoughsome researchers have observed a surrounding membrane (31, 56, 72), it is generallyconsidered to be absent (46, 72, 114).

The presence of other constituents in the granules, e.g. RNA, DNA, protein, andlipid, is still in doubt (31, 46, 82, 139). Neimeyer & Richter (82) found that polyphosphate of ~4nacystis nidulans was extracted with the nucleic acids but couldbe separated by th!n-layer and column chromatography. The granules were recentlyisolated from Micrococcus lysodeikticus and found to contain 24% protein, 30%lipids, and 27% polyphosphate; some carbohydrate, nucleic acid, and polyvalentcations were also detected (31). The M. lysodeikticus granules do have an unusualappearance, but it appears that this preparation is grossly contaminated. Earlierreports (46) indicate that the granules are composed primarily of polyphosphatewith only trace amounts of DNA, RNA, and protein.

The granules occur at a fairly constant number during the exponential phase ofgrowth when phosphate is in excess (23, 31,138, 139), and decrease under limitingphosphate conditions (82, 139) or during periods of cell inactivation (23, 31).

The polymer is synthesized by the enzyme, polyphosphate kinase, which catalyzesthe following reaction: ATP + (HPO3)n ~ ADP + (HPO3)n + ~ (46). HOW thepolymer is organized into a granule is not known, but Voelz et al (129) proposedthat the polymer strands are precipitated onto cytoplasmic material and becometightly coiled into granules. Harold (45) suggests that the polymer is precipitatedby the high ionic strength of the cytoplasm. Jensen (57) reports a number of stagesin granule development in blue-green algae including the formation of an electrontranslucent area in the cytoplasm, a porous body, and finally the mature, electron-opaque granule. What these stages represent must await critical isolation and char-acterization studies.

The polymer can be degraded by a number of enzymes (46): polyphosphatekinase, polyphosphate-adenosine monophosphate-phosphotransferase, polyphos-phate glucokinase, polyphosphate fructokinase, and polyphosphatases.

The polyphosphate may function as a phos.phagen, i.e. energy storage, as a phos-phate reserve, or as a regulator of phosphate economy (46). Its primary functionappears to be phosphate storage, but its possible role in the storage of energy shouldnot be overlooked (8, 23, 47, 124).

Cyanophycin (Structured) Granules

Cyanophycin granules are observed in most, if not all, of the blue-green algae (72).When well preserved by proper fixation, they appear in thin sections (Figure 2)


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INCLUSION BODIES OF PROKARYOTES 171

"tightly packed, undulating, flattened sacs" (72). These inclusions are without limiting membrane and are variable in size and shape; diameters of greater than 500nm have been reported in older cells (72, 73). Simon (115) isolated the granules Anabaena cylindrica and found them to consist of 25,000-100,000 dalton molecularweight polypeptides containing arganine and aspartic acid in a 1:1 ratio.

The number of granules is lowest in cells of exponentially growing cultures andhighest in cells of stationary phase cultures and akinetes. The number decreasesupon germination of the akinete or upon transfer of the maximum stationary phaseculture to conditions suitable for growth (72, 78, 116). Furthermore, as the cyano-phycin granule polypeptide increases in the late exponential phase of growth,phycocyanin decreases.

Addition of chloramphenicol to exponentially growing cultures of A. cylindricaresulted in a rapid cessation of protein synthesis and growth, but an accumulationof cyanophycin granule polypeptide; the polypeptide is therefore not synthesized bya mRNA-ribosomal system (117). Upon removal of the chloramphenicol, growthresumes and the cyanophycin granule polypeptide is hydrolyzed. The cyanophycingranule polypeptide-synthesizing enzymes must be present in the cells of exponen-tially growing cultures; chloramphenicol rapidly inhibits protein synthesis and thepolypeptide is then produced. The synthesis is an energy dependent process; (3,4-dichlorophenyl)-l,l-dimethyl urea, a specific photophosphorylation inhibitor,blocks the synthesis of the polypeptide after the addition of chloramphenicol (117).

All of the information is consistent with the theory that the granules are cellularnitrogen reserves (117). During normal growth, amino acids are incorporated intoprotein, but upon cessation of growth, nitrogen fixed from the atmosphere is storedin the polypeptide.

The observation (lg) that the cyanophyein granules participate in the synthesisof polyphosphate needs to be carefully evaluated.

Phycobilisomes (10, 139)

Three biliproteins have been reported in the blue-green algae: C-phycocyanin(C-PC), allophycocyanin (allo-PC), and C-phycoerythrin (C-PE). C-PC and PC appear to be universal constituents; C-PE may or may not be present. Thechromophores, the tetrapyrrole pigments phycocyanobilin and phycoerythrobilin,are readily separated from their apoproteins (10, 139). The C-PC, allo-PC, and C-PEhave maximum absorbance peaks at 615 nm, 650 nm (shoulder at 620 nm), and 565nm, respectively. The molecular weights of the biliproteins have been a subject ofsome controversy. The most recent evidence indicates that the monomers of C-PCand C-PE have molecular weights of ca 30,000 and 40,000 daltons, respectively (10,66, 125). Furthermore, the monomers, in both instances, are composed of twopolypeptides of unequal size (66, 125, 139). Because the monomers aggregate particles of varying sizes it is difficult to ascertain what the predominant in vivospecies is for each biliprotein; a hexameric structure appears to be most likely (10).

The biliproteins constitute 14-28% of the call dry weight, depending on theorganism examined and growth conditions, and function in the harvest of lightenergy and its subsequent transfer to chlorophyl (37, 139).


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172 SHIVELY

Phycobilisomes, high molecular weight aggregates of C-PC, allo-PC, and fre-quently C-PE, were originally demonstrated in the Rhodophyta, but have subse-quently been observed in many blue-green algae (22, 27, 37, 75, 136, 137). Theseinclusions (Figure 5) are 35-50 nm in diameter and are attached to the photosyn-thetic lamellae (22, 27, 37, 75, 139). It is still to be resolved whether all of thebiliproteins are attached, or ifa certain quantity is free in the cytoplasm (22, 27, 37,139).

The C-PC containing phycobilisomes of Synechococcus lividus are described asrods, 35 nm in diameter, consisting of heptamers of smaller rods each of which iscomposed of a stack of dimeric discs (22). Each dimeric disc is 3.0-3.5 nm thickand 12.0-12.5 nm in diameter. It is proposed that these discs are hexamers ofphycocyanin. The phycobilisomes of Anacystis nidulans also lack C-PE and are rodshaped (37), while those of Nostoc species (Figure 6), which possess all threebiliproteins, are compact structures with a rounded surface and a flattened base (40nm in diameter) which attaches the complex to the photosynthetic lamellae (37).

Isolated phycobilisomes (Figure 7)are variable in size and shape (37). This be the result of natural heterogeneity or damage during isolation.

Crystals and Paracrystalline Arrays

Several members of the genus Bacillus form parasporal inclusions (79, 84, 101). Thebest characterized, because of its toxicity to Lepidoptera larvae, is the parasporalcrystal of Bacillus thuringiensis (84). The crystal (Figure 8), a bipyrimidal octahedrawith a square base plane, is composed of rod-shaped subunits 4.7 nm by 11.8 nm.Norris (84) reports that the rod-shaped subunits have a molecular weight of 230,000daltons and break down to smaller subunits upon treatment with alkali. Whetherthere is one or more polypeptide species is a controversial subject which has beenaggravated by the use.of different strains and solubilizing agents. Sayles et al (107)reported the existence of a heterogenous population of small polypeptides withmolecular weights of 1,000-1,500 daltons. Herbert et al (49) found two subunitswith molecular weights of 55,000 and 120,000 daltons in a ratio of 2 to 1 and Glatronet al (35) discovered only a single subunit of 80,000 dalton mol wt. Thecontroversy is further deepened by the discovery that three antigens are associatedwith the spore crystal (74, 84). Norris (84) attributes these to different subunits, Lecadet & Dedonder (74) believe that they may result from the association of identical subunit. The crystal protein or proteins are synthesized by RNA-mediated

Figure 5 Phycobilisomes (arrows) in Microcoleus vaginatus. Courtesy of R. B. Wildman(136); with permission.

Figure 6 Phycobilisomes (arrows) in Nostoc species. Courtesy orE. Gantt (37); with permis-sion.

Figure 7 Isolated phycobilisomes of Nostoc species: ~negative stain. Courtesy of E. Gantt,Smithsonian Institution, Radiation Biology Laboratory, Rockville, Md.

Figure 8 Freeze-etching of the parasporal crystal of Bacillus thuringiensis vat. alesti. Cour-tesy of S. C. Holt, University of Massachusetts, Amherst.


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protein synthesis (142) and crystal formation may be initiated on the spore exo-sporium (118), or on cytoplasmic membrane invaginations (100).

The formation of the crystal is definitely linked to sporulation (84, 100, 142) andevidence indicates that the crystal protein may result from the over production ofspore coat protein(s) (74, 119). In this regard, Santo & Doi (106) report crystalformation in a RNA polymerase mutant of Bacillus subtilis which is conditionallytemperature sensitive only during sporulation (Figure 9). The crystal forms in 1-2%of the cells that do not form spores (85% of calls sporulate normally). The crystalsubunits show a center to center repeat distance of 8.4 nm; the cross striation unitsaveraged 8.5 nm by 8.5 nm. This material may be a spore protein.

Parasporal crystals have also been reported in clostridia. Pope et al (95) discov-ered a crystal lattice (250 nm by 160 nm by 130 rim) composed of rods (12 diameter) with a repeat distance of 15 nm in Clostridium cochlearium (Figure 10).This crystal was formed only in the sporangium. A second crystal was formed (6.5nm periodicity) in both vegetative cells and sporangia. Norris (85) speculates thatthese crystals are of degenerate bacteriophage protein(s) which he has also noted B. thuringiensis. More recently (20), parasporal paracrystalline inclusions (192 by 2,120 nm) have been found in Clostridium perfringens (Figure 11). They areformed only during spore morphogenesis and only by enterotoxin positive strains.It is theorized that the crystals may represent the enterotoxin.

An interesting intraeytoplasmic axial fiber with a terminal structure was recentlyreported in a variant (rho) form of Mycoplasma (89). The fiber (40-120 nm wide)which extends the length of the cell (Figure 12) is composed of alternating dark andlight bands with a banding periodicity of 12.0-14.5 nm (Figure 13). The structureis apparently composed of fibers, laying parallel to the long axis. More recently (103)it has been shown that the fiber is a paracrystalline aggregate of fibrous protein(molecular weight 28,000 daltons) which can be disassembled and reassembled intothe fiber; small amounts of other proteins may also be present.

Kim & Barksdale (65) reported the presence of a crystal in 10% of the cells old cultures of bacterium 22 M (Figure 14). They consist of parallel rows of 7.5 in diameter polyhedral subunits, 2.0 nm apart and 3.0 nm between rows. Theseoccur in cells showing degenerative processes and are not likely to be of viral origin(65).

Figure 9 Crystal of RNA polymerase mutant of Bacillus subtilis. Courtesy of L. Santo (106);with permission.Figure 10 Parasporal crystal of Clostridium cochlearium. Courtesy of L. Pope (95); withpermission.Figure 11 Paracrystalline inclusion in mutant 8~, of Clostridium perfringens. Symbols: I,inclusion; OC, outer spore coat material; IC, inner spore coat material. Courtesy of C. L.Duncan (20); with permission.Figure 12 Axial fiber in a variant (rho) form of Mycoplasma: negative stain. Courtesy of J.E. Peterson (89); with permission.Figure 13 Segment of an isolated axial fiber of a variant (rho) form of Mycoplasma: negativestain. Marker bar equals 200 nm. Courtesy of J. E. Peterson (89); with permission.

Figure 14 Crystal in bacterium 22 M. Courtesy of L Barksdale (65); with permission.


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Crystalline inclusions have also been observed in the blue,green algae (6, 32, 59),in a root nodule endophyte (33), in the symbiotic bacteria of a leafhopper (67), in Thiobacillus intermedius (113).

Tubules, Microtubules, and Raphidosomes

These inclusions have been observed in a number of bacteria (12, 17, 55, 69, 88, 94,97, 126, 140). They are variable in length in different organisms and sometimes inthe same organism. Diameters from 9,0-40.0 nm have been reported (17, 55, 94,126). Those of Saprospira grandis contain an outer sheath 23.0 nm in diameter (17).Some are formed only during cell lysis, are composed of lipid and protein, and areproposed to be reorganized mesosome or cytoplasmic membrane (69, 88, 126).Others are thought to be degenerative bacteriophage proteins (12~ 17, 97). If someof these structures are shown not to be the result of a degenerative bacteriophagegenome, their cellular function could prove interesting.

MEMBRANE-ENCLOSED INCLUSIONS

The barrier surrounding all of these inclusions is of the nonunit membrane type, i.e.in thin section it appears as a single layer, 2.0-4.0 nm thick. Current evidence (poly-fl-hydroxybutyrate, sulfur globules, gas vacuoles) indicates that the membrane iscomposed entirely of protein.

Polyglucoside (Glycogen) Granules

Many prokaryotes have been shown to accumulate polyglucose; however, in mostinstances the polymer accumulates as a nonmembrane bound granule (see above).In several species of Clostridium [CI. botulinum type E (5), CI. butyricum (104),CI. pasteurianum (71, 76), and CI. saccharobutyricum (104)], the polyglucose gran-ules appear to be surrounded by a single-layered membrane (Figure 15). Membrane-bound polyglucose granules have not been observed in Clostridium tyrobutyricum(105) and CI. botulinum type A (5).

The granules first appear in the cells of early log phase cultures and become morenumerous as the culture ages, reaching a maximum (ca 15% of cell dry weight) the outset of sporulation (5, 71, 76, 122). Their initial appearance correlates wellwith the derepression of ADP-glucose pyrophosphorylase (71). The polymer ap-pears to be of the amylopectin type (5, 34, 122) and molecular weights of 180,000daltons have been recorded. Laishley et al (71) recently reported on the isolationand morphology of the granules (160-300 nm in diameter) of CI. pasteurianum.

Figure 15 Polyglucoside granules in Clostridium pasteurianum. Courtesy of E. J. Laishley(71). (Reproduced with permission of the National Research Council of Canada.)

Figure 16 Poly-/3-hydroxybutyrate granules Thiobacillus (Ferrobacillus) ferrooxidans. Cour-tesy of D. G. Lundgren, with permission (75a).

Figure 17 Sulfur globules in Chromatium weissii. Marker bar equals 1 /xm. Courtesy of G.L. Hageage, National Institute of Dental Research, Bethesda, Md.


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Freeze-etch studies revealed 12.0-16.0 nm in diameter projections in the granulematrix; these projections were hypothesized to be polymer fibrils.

Evidence indicates that the enzyme, granulose synthetase, is intimately associatedwith the granules (102). One could speculate that the enzyme may be an integralpart of the granule membrane.

Poly-/7-hydroxybutyrate Granules (19)

The ability to accumulate poly-fl-hydroxybutyrate (PHB) is common among thebacteria (19, 26, 48, 93, 121,123, 133, 141) and has been reported in the blue-greenalgae (60, 61). PHB has recently been found to accumulate in sporulating cells CI, botulinum type E (26). As already noted, this organism has also been shown deposit polyglucoside granules (5, 122).

The polymer accumulates, sometimes to greater than 50% of the cell dry weight(19), when carbon and energy sources are in excess. PHB is considered to be cellular reserve of energy, or of carbon and energy (19).

The PHB is deposited in nonunit membrane-enclosed granules (Figure 16) whichappear electron transparent in thin section (61,133). Granule diameters of 100-800nm (21, 25, 60, 141) have been reported and each granule may contain severalthousand PHB molecules (25). The granules consist of 98% PHB, 2% protein, andtrace amounts of lipid and phosphorous (19, 39). The surrounding membrane 2.0-4.0 nm thick (2, 60, 133) and may arise from one layer of the cytoplasmicmembrane (91). Dunlop & Robards (21), using freeze-etching, have recently demon-strated that the granules (240-720 nm in diameter) of Bacillus cereus consist of acentral core (140-370 nm in diameter) which occupies less than 50% of the granulevolume, an outer coat, and a surrounding membrane. This may indicate that thepolymer is in different physical states within the granule.

The molecular weights of PHB vary to a great degree; values from 1,000 to256,000 daitons have been reported (19). The variation may be the result of prepara-tive techniques and/or organismic and growth state differences. Research indicatesthat the PHB in the granule is in a crystalline form (19, 25). The crystallineconformation 9f the polymer is a right-hand helix stabilized by carbonyl-methylinteraction (19).

The enzyme for polymerization of o-(-)-fl-hydroxybutyryl CoA, PHB synthe-tase, and all or part of the depolymerizing complex (depending on the organism)are associated with the granules, presumably as part of the surrounding membrane(19, 38, 39). It is postulated that the PHB synthetase aggregates into a mieellar formand the PHB is deposited within (25). The reutilization of the PHB, under condi-tions of energy and/or carbon starvation, involves a labile, protein, inhibitor factor(commonly associated with the granules); and activator, which counteracts theeffect of the inhibitor; and the depolymerase (19, 38). The activator and depolyme-rase may or may not be associated with the granules (19). Barber & Nakata (2) recently demonstrated that the initial step in the reutilization of PHB in B. cereusis the rupture of the granule membrane; a heat labile, trypsin sensitive protein wasisolated. How this factor is involved in the above depolymerization complex remains


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INCLUSION BODIES OF PROKARYOTES179

to be elucidated. Monomers and dimers of fl-hydroxybutyrate are the resultingproducts; the dimer is further hydrolyzed by dimer hydrolyase (19).

Sulfur Globules

Sulfur globules are a distinctive feature of the Thiorhodaceae and of certain otherapochlorotic sulfur bacteria (16, 19, 29, 40, 77, 99). The sulfur is deposited as thecells oxidize and grow in the presence of hydrogen sulfide. The sulfur globulesdisappear, i.e. the sulfur is oxidized, when hydrogen sulfide becomes limiting (77).

The globules are generally observed in thin sections as nonunit memt, rane-enclosed holes; the sulfur is removed during dehydration (Figure 17). They varyfrom 100 nm to over 1.0 p,m in diameter and are most commonly deposited ininvaginated pockets of the cytoplasmic membrane, i.e. they are inside the cell wall,but outside the cytoplasmic membrane (40, 77). Therefore, it may be erroneous classify the sulfur globules as inclusions. However, the sulfur globules of Thiovulummajus (16, 29) and, in some instances, those of Beggiatoa and Chromatium (G. J.Hageage, personal communication) are found in the cytoplasm proper.

The globules have a monolayer membrane (16, 29, 40, 81, 99, 108) which consistsof globular subunits, 2.5 nm in diameter (81), and is composed entirely of protein,molecular weight 13,500 daltons (109). The protein, solubilized by urea, reaggre-gated into sheets when the urea was removed by dialysis (109). $chmidt et al (109)speculated that the membrane provides binding sites for the enzymes responsible forsulfur metabolism.

Polarizing microscopy and X-ray diffraction studies on globules in intact ceils orisolated (wet state) revealed the sulfur to be in liquid state (40, 41). During dryingthe sulfur passed through an unstable crystalline phase and was eventually con-verted to crystalline orthorhombic sulfur (40, 41).

Gas Vacuoles (131, 132)

Gas vacuoles occur in many aquatic prokaryotic organisms including representa-tives of the blue-green algae, the photosynthetic green and purple sulfur bacteria,and a few other bacteria (Figures 2 and 18).

The gas vacuoles are complex organelles consisting of an array of substructuresreferred to as gas vesicles. The gas vesicles are hollow cylinders with conical ends(Figure 19). The diameter and length vary from 65-115 nm and 0.2-1.2 ~m,respectively. The limiting barrier of the gas vesicle varies between 1.9 and 3.0 nmdepending on the organism and the method of examination employed (thin-section-ing, shadow casting, X-ray diffraction). The vesicles have striated fibs, with periodicity of 4.0-5.0 nm (depends on method of examination), at a fight angle the long axis of the vesicle (Figure 19). Therefore, the vesicle is proposed to composed of either a stack of hoops or one to several continuous strands. It has beensuggested that the vesicles are built of subunits 2.8-3.5 nm in diameter. This conclu-sion is supported by the observation of a beaded appearance in thin section whenthe vesicles are cut longitudinally at a right angle to the plane of the striated ribs.


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X-ray diffraction studies also indicate subunits, but whether these are the same asthose observed in electron microscopy remains to be proven.

The membrane appears to consist entirely of protein; phosphorous and galactosemay occur in some species (68). The vesicles of the blue-green algae are partiallysoluble in 80% formic acid or phenol-acetic acid water and consist of a single proteinspecies, 14,000-15,000 daltons mol wt. The gas vesicles of Halobacterium halobiumare not soluble in the above reagents or guanidine hydroehloride, organic solvents,and pH extremes (68). Neither the blue-green alga or Halobacterium vesicles aresoluble in detergents (28, 68).

The pHI of the blue-green alga vesicle protein is 7.0 and that of H. halobium is4.0. The low pHI may reflect the phosphorous content of these vesicles (68). All the vesicles thus far isolated have a predominance (30-33%) of amino acids withhydrophobic side chains. It has been hypothesized that the inner surface of thevesicle membrane is hydrophobic; the outer surface hydrophilic.

The membrane is rigid, i.e. it is not inflated with gas, impermeable to water, andfreely permeable to all gasses. Its impermeability to water is attributed to thehydrophobic inner membrane surface. The vesicle appears to be a self-assemblingstructure; the body elongates after formation of the conical ends. The shape isdetermined by the subunit proteins.

The g~s vacuoles may function in buoyancy provision, buoyancy regulation, lightshielding, surface to volume regulation, or a combination of these functions.

Polyhedral Bodies (Carboxysomes)

Inclusions with polygonal profiles (Figure 20) have been observed in the blue-greenalgae (32, 51, 59, 87, 139), and in many, but not all, of the nitrifying bacteria (80,93, 127, 134) and thiobacilli (113, 133). They have also been observed Beggiatoa(D. S. Hoare, personal communication). There are commonly several bodies per cell;t-15 (4-6 most common) in Thiobacillus neapolitanus (112), 60-70 in older cellsof Thiobacillus thioparus (113), and 200 in Nitrococcus mobilis (134)..

As seen in thin section, the bodies are 90-500 nm in diameter (32, 51, 87, 112,127, 134), have a granular substructure of medium electron density (87, 112, 139),are commonly located in the nucleoplasmic region of the cell (87, 112, 113), andappear to be bounded by a nonunit membrane 3.0-4.0 nm thick (87, 112, 134). tripartite membrane less than 5.5 nm thick has been observed in some blue-greenalgae preparations (135).

The inclusions have been recently isolated from T. neapolitanus (111). By elec-tron microscopy, the purified bodies (Figure 21) were shown to consist of a mass

Figure 18 Freeze-etching of Nostoc rnuscorum showing gas vacuoles. Courtesy of J. R.Waaland, University of Washington, Seattle.

Figure 19 Isolated gas vesicle from Anabaenaflos-aquae: negative stain. Courtesy of A. E.Walsby (131); with permission.

Figure 20 Polyhedral bodies (carboxysomes) in Thiobacillus neapolitanus J. M. Shively(113); with permission.


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of 10 nm particles in a paracrystalline array surrounded by a membrane 3.5 nm thick(111, 112). The 10 nm particles (Figure 22) have been identified as ribulose-l,5-di-phosphate carboxylase (111). Carboxysomes is the proposed name for the polyhe-dral bodies (111), The bodies are fragile and current assay conditions do not favortheir stability. The presence.of other components, e.g. other enzymes, remains tobe established.

That the bodies from all the organisms are identical is still in doubt, but circum-stantial evidence strongly supports this conclusion. All of the organisms utilizecarbon dioxide as their sole source of carbon; in thin section the bodies all haveessentially the same appearance; and negative stains of released but impure bodiesof Anacystis nidulans (32) and N. mobilis (134) show a membrane-enclosed, para-crystalline array of particles. Research is in progress in this laboratory to prove thehypothesis.

It is tempting to propose that the bodies are active in carbon dioxide fixation, butalternatively they may be for enzyme storage. Bock& Heinrich (4) observed greaternumbers of polyhedral bodies in cells of Nitrobacter winogradskyi stored for 200-300 days in the absence of nitrite than in reactivated cells; the loss of the bodiescorrelated with the degree of reactivation. Furthermore, as the reactivation pro-ceeded the cell membranes became studded with particles resembling ribulose-1,5-all-phosphate carboxylase. They also observed that 37% of,the reactivated cells didnot possess bodies. Remsen & Lundgren (98) published a micrograph ofa Thiobacil-lus (Ferrobacillus)ferrooxidans cell which did not possess polyhedral bodies. If theyare essential entities, why are they absent in some cells? It is also well known thatin the blue-green algae, ribulose-l,5-diphosphate carboxylase synthesis is not re-pressed even in the presence of organic substrates, i.e. cultures grown in the darkwith an organic substrate do not have reduced enzyme levels (7). Dark grown cells,even in the absence of carbon dioxide, have polyhedral bodies (51). One couldtheorize then, that since the cells do not have the ability to control ribulose-l,5-di-phosphate carboxylase synthesis, the enzyme is packed away in a nonactive stateuntil needed. The present evidence then favors a storage function, but criticalexperimentation needs to be undertaken. No information is available on how thebodies are assembled or disassembled.

Chlorobium Vesicles

The chlorophyl, principally chlorobium chlorophyl, but also bacteriochlorophyl a,of the green photosynthetic bacteria is contained within vesicles (Figure 23) 30-40nm wide by 100-150 nm long, which are bounded by a nonunit menbrane 2-3 nm

Figure 21 Isolated carboxysomes of Thiobacillus neapolitanus." negative stain.

Figure 22 Ribulose-l,5-diphosphate carboxylase from carboxysomes of T. neapolitanus:negative stain.

Figure 23 Chlorobium vesicles in a ballistically opened cell of Chloropseudomonas ethylicum:negative stain. Courtesy of S. C. Holt (53); with permission.


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184 SHIVELY

thick (13,. 15, 52, 90). The vesicles immediately underly the cytoplasmic membraneand more or less completely line the internal periphery of the cell (13, 52, 53). Thevesicles may appear either transparent or opaque in thin section depending on theembedding medium utilized (13). The vesicles themselves may be interconnected(52), but they have little, if any, association with the cytoplasmic membrane of thecell (I 5). Under normalgrowth conditions the vesicles constitute about 12% of thetotal cell dry weight (15); their number and their chlorophyl content is inverselyproportional to the light intensity at which the cells are grown (53).

Some substructure has been observed. Cohen-Bazire et al (13) reported that thevesicles were filled with 1.0-2.0 nm wide fibrils laying parallel to the long axis of thevesicles. Circular particles, 9.0-10.0 nm in diameter, with a center hole and consist-ing of 5-6 subunits have also been seen (15).

On a dry weight basis, the vesicles are composed of 50% lipid, 30% protein, and15% carbohydrate (15). Seventy-five percent of the vesicular lipid is in the chlo-robium chlorophyl; the remaining 25% is glycolipid I, monogalactosyl diglyceride.Glycolipid I is not found in the cytoplasmic membrane of the cell where anotherglyeolipid, glycolipid II, and all of the phospholipids reside. The vesicles haveNADH- and NADPH-linked dye reductases, as do the cytoplasmic membranes, butlack the cytoplasmic membrane-associated enzymes, malic and succinic dehydroge-nase (15). Fowler, et al (30) recently isolated a bacteriochlorophyl a reaction centercomplex from the vesicles of Chlorobium limicola and Chlorobiurn thiosulfato.philum. The polymeric complex with a minimum molecular weight of 1.5 X 104daltons is composed of similar subunits and co~atains bacteriochlorophyl a, bchl-aprotein, cytochrome 553, carotenoids, chlorophyl P-340, a small amount of chlo-robium chlorophyl, and possibly other components. In this complex bacterio-chlorophyl a alone absorbs light energy.

Obviously, the green photosynthetic bacteria have a much greater degree ofstructural and functional differentiation than do the other photosynthetic bacteria(90) or, for that matter, the blue-green algae (139).

OTHER INCLUSIONS

Lipid deposits have been repeatedly reported in a number of bacteria and blue-greenalgae (138, 139). Many of these will probably be identified as poly-fl-hydroxybuty-rate, but the occurrence of other lipid deposits is entirely possible. Other inclusionsin prokaryotes include the cylindrical bodies of Trichodesmium erythraeum (1) andSymploca muscorum (87); the membrane-associated protein inclusions of Bacillussubtilis (3); the striated organelles of Halobacterium halobium (11); the fibrillarorganelle (16) and greenish bodies (50) Thiovulum majus," thefila mentous inclu-sions, spheroids, rosette-like inclusions, and intrathylakoidal granules of representa-tives of the blue-green algae (59); the large granulated body of Corynebacterium(63); the hydrocarbon inclusion of dcinetobacter (64); the calcium carbonate inclu-sions of A ch roma tiu m (120); and the ribosome studded inclusions of the my xosporesof Stigmatella arthantiaca (128).


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CONCLUDING REMARKS

A wealth of information is available on the inclusion bodies of prokaryotes, but itis obvious that much remains to be revealed. With the availability of sophisticatedexperimental techniques, it is time to completely elucidate the structure of theinclusions, study the mechanisms of their assembly and disassembly, and fully assesstheir metabolic roles in the prokaryotic organisms.

ACKNOWLEDGMENTS

I am deeply indebted to those who provided micrographs; .to those who suppliedprepublication information; to Clanton Black, University of Georgia, Athens, forhis assistance with the literature sui’vey; and to Sue Bogardus, Oya Yazar, and H.W. Raynall for their assistance in preparing the manuscript.

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inclusion bodies of prokaryotes

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