The text below contains more information about Bacteroides than you ever knew existed. It was taken from the introduction of Gena Tribbles' Doctoral thesis entitled: Development of a Model of Transposition for the Bacteroides mobilizable transposon TN4555.

Anaerobes comprise the majority of bacteria in the human colon; the most numerically predominant of these are members of the genus Bacteroides. Originally described in 1898 (115), for many years the Bacteroides were a vague conglomeration of host-associated, obligately anaerobic, gram-negative, pleomorphic rods that could not be convincingly assigned to any other genera. Physiological analysis of this genus revealed considerable heterogeneity with regard to their biochemical properties, indicating these bacteria did not represent a true phylogenetic grouping. With the advent of phylogenetic analysis techniques, several investigators have tried to redefine this group of bacteria using physiological characteristics (39), serotyping (50), bacteriophage typing (6), lipid analysis (59), oligonucleotide cataloging (73), and 5S - 16S rRNA sequence comparisons (46, 72, 113, 116 ). Based on this information, the original Bacteroides members have been partitioned into three genera: Bacteroides (99), Prevotella (97), and Porphyromonas (98). The Bacteroides are found predominantly in the colon of mammals, while the Prevotella and Porphyromonads generally are associated with the oral cavity and the rumen.
The current definition of Bacteroides species is as follows: a) obligately anaerobic, Gram-negative, b) saccharolytic, producing acetate and succinate as the major metabolic end products, c) contain enzymes of the hexose monophosphate shunt-pentose phosphate pathway, d) have a DNA-base composition in the range 40-48 mol% GC, e) membranes contain sphingolipids, and contain a mixture of long-chain fatty acids, mainly straight chain saturated, anteiso-methyl, and iso-methyl branched acids, f) possess menaquiones with MK-10 and MK-11 as the major components, and g) contain meso-diaminopimelic acid in their peptidoglycan (96). This definition restricts the Bacteroides to ten species: B. fragilis, B. thetaiotaomicron, B. vulgatus, B. ovatus, B. distasonis, B. uniformis, B. stercoris, B. eggerthii, B. merdae, and B. caccae, with B. fragilis as the type strain. The Bacteroides, along with Prevotella and Porphyromonas, form one major subgroup in the bacterial phylum Cytophaga-Flavobacter-Bacteroides (27). This phylum diverged quite early in the evolutionary lineage of bacteria, and thus the Bacteroides, although gram-negative organisms, are not closely related to the enteric gram-negatives such as Escherichia coli (Fig. 1.1).

FIG. 1.1. Phylogenetic tree of Eubacteria based on 16s rRNA sequence comparisons. The evolutionary relationship between prokaryotic phyla are shown. Branch lengths on the tree represent evolutionary distance. Grey wedges represent divergence within the individual phyla. Derived from (116). (Click image for larger version)

The Bacteroides inhabit the human colon, which contains the largest, most complex bacterial population of any colonized area of the human body. The colonic contents contain in excess of 1011 organisms per gram of wet weight, representing over 400 species (61). The Bacteroides are the most numerous members of the normal flora, representing nearly 1011 organisms per gram of feces (dry weight) (25). Gut organisms are involved in numerous metabolic activities in the colon, including fermentation of carbohydrates, utilization of nitrogenous substances, and biotransformation of bile acids and other steroids (36). In order to maintain their high numbers, the Bacteroides are evidently able to compete with other members of the flora, as well as transient organisms, for utilization of these resources. While the role of the microflora in the physiology of the human intestine is not well studied, it is clear that the anaerobic members of this ecosystem play a fundamental role in the processing of complex molecules into simpler compounds, and through their metabolic activities the human microflora participate in the complex physiology of the host (8).
Most intestinal bacteria are saccharolytic, obtaining carbon and energy by hydrolysis of host and dietary carbohydrate molecules (75). Simple sugars are rarely encountered in the colon as most are absorbed in the small intestine, however it is estimated that approximately 2% of simple sugars can pass through the upper gastrointestinal tract when large amounts of starch and complex carbohydrates are also present during digestion (92). Bacteroides species are able to utilize simple sugars when present (44, 95), but due to their limited availability, simple sugars are probably not the main source of energy for the Bacteroides. Much more prevalent in the colon are polysaccharides, from dietary sources and host cells. Polysaccharides from plant fibers, such as cellulose, xylan, arabinogalactan, and pectin, and vegetable starches such as amylose and amylopectin contain Bacteroides have been shown to have a variety of glucosidase activities, including a b-1,3-glucosidase activity responsible for laminarin degradation (94), and a variety of a and b-1, 4 and -1, 6 xylosidase and glucosidase activities induced by the presence of hemicellulose (83). Originally it was believed that these enzymatic activities were extracellular, and the short oligosaccharides and monosaccharides produced by hydrolysis were taken up into the cell for fermentation. Analysis of the B. thetaiotaomicron starch utilization system (sus) (16), has revealed the polysaccharides to be bound to an outer membrane receptor system (85), and pulled into the periplasm for degradation into monosaccharides. The Bacteroides use a similar approach for uptake and degradation of chondroitin sulfate (10), indicating this technique may provide a competitive advantage in the human gut, as polysaccharides sequestered in the periplasm are less likely to be "stolen" by other intestinal organisms or lost by diffusion.
Interestingly, utilization of chondroitin sulfate by Bacteroides thetaiotaomicron is repressed in the presence of glucose (93), while utilization of other sugars in B. thetaiotaomicron is tightly regulated in the presence of mannose (52). This implies the Bacteroides may have a catabolite repression mechanism to allow for the utilization of select carbon sources in preference to others. If so, this system is probably not similar to the catabolite repression systems of enteric bacteria, as the Bacteroides do not possess cyclic AMP (42). It is likely that most Bacteroides polysaccharide utilization systems are controlled by repressor/inducer mechanisms, as B. ovatus and B. thetaiotaomicron are able to utilize several sugars simultaneously (52), and several polysaccharide utilization genes have been shown to be activated in the presence of their substrate (18, 120, 81).
Carbohydrate fermentation by the Bacteroides and other intestinal bacteria results in the production of a pool of volatile fatty acids, predominately acetate, propionate (from succinate), and butyrate. These short chain fatty acids are reabsorbed through the large intestine, and utilized by the host as an energy source (56). It has been estimated that absorption of the short chain fatty acids could provide up to 540 kcal/d, a significant proportion of the host's daily energy requirement (15).
The utilization of nitrogen sources by the intestinal Bacteroides is not well understood, as most work in the area of nitrogen uptake has been done with rumen organisms. However, several parallels may be drawn between intestinal and rumen bacteria, providing a paradigm of nitrogen utilization in the human gut. There are three major sources of nitrogen in the mammalian intestine: dietary protein, epithelial cell and mucin glycoproteins, and ammonia (38). Most dietary protein is degraded and absorbed before reaching the large intestine, but once in the colon, these peptides and amino acids are not able to be absorbed by the host (122). Instead, a two step degradation process occurs, during which peptides are proteolysed to amino acids, which are subsequently deaminated to form ammonia, CO2, volatile fatty acids, and branched chain fatty acids (122). The ammonia is utilized by the intestinal bacteria as a nitrogen source (38). Bacteroides fragilis has been shown to produce three major proteases (28), with activity against a variety of proteins, including casein, trypsin, and chymotrypsin, but not collagen, elastin, or gelatin (29). The Bacteroides also encode glutamine synthetase (108) and glutamate dehydrogenase (4), which are important for ammonia assimilation but the regulation of these activities is not yet understood.
The Bacteroides play a key role in the enterohepatic circulation of bile acids. Cholic acid and chenodeoxycholic acid are the two main bile acids synthesized in the human liver, where they are conjugated to taurine or glycine polar side groups before secretion in bile. Once bile enters the gut, the conjugated bile acids assist in the absorption of dietary fats. If the bile acids are not reabsorbed in association with fat in the upper intestine, they are deconjugated by bacteria to secondary bile acids, primarily deoxycholic and lithocholic acid, although the microflora can generate 15-20 other secondary bile acids from these same precursors (41). Deconjugation allows the bile acids to reenter the enterohepatic circulation via the portal system, where they are returned to the liver and reconjugated for further use (5). The secondary bile acids deoxycholic and lithocholic acid are produced by 7 alpha-dehydrogenation of the primary bile acids (41); once these secondary bile acids are produced, a variety of other bacterial reactions can occur, including oxidation-reduction, desulphation, and dehydrogenation (24), producing a variety of isomers of secondary bile acids. The Bacteroides have been found to play a major role in the biotransformation of bile acids, and contain many enzymes required for these reactions, including a hydrolase (109), dehydrogenase (43), and dehydroxylase (21, 87). The direct benefit to the host is obvious, as deconjugation of the primary bile acids allow them to be reabsorbed in the large intestine instead of lost in the feces. The benefit to the Bacteroides and other intestinal bacteria is not clear, but may contribute to energy metabolism.
Aside from their metabolic activities, the Bacteroides and other anaerobes provide an additional benefit to their host in excluding pathogenic organisms from colonizing the intestine (114). Colonization resistance mediated by anaerobes is thought to occur by four mechanisms: competition for nutrients, competition for intestinal wall attachment sites, production of volatile fatty acids, and release of free bile acids (37). The intestinal microflora adhere to the surface of epithelial cells and mucin associated with the intestinal wall, with Bacteroides being the most common anaerobic colonizer (11). By coating the walls of the intestine, it is believed that the microflora prevent transient bacteria from obtaining a binding site on the intestinal surface, and the transients are subsequently lost with the luminal contents during peristalsis. The volatile fatty acids produced as metabolic end products by the Bacteroides are also believed to play a role in colonization resistance, by lowering the pH and oxidation-reduction potential of the intestinal milieu, resulting in unfavorable growth conditions for transient bacteria (37). The most notable pathogens inhibited under these conditions are Salmonella enteritidis (57), and Shigella flexineri (53). Production of free bile acids also plays a role in inhibition of pathogens, as bile salts are toxic to many organisms, including Clostridium botulinum (40).

While the Bacteroides occupy a significant position in the normal flora, they also are opportunistic pathogens, primarily in infections of the peritoneal cavity. B. fragilis is the most notable pathogen; although it makes up only 1-2% of the normal flora, it is the Bacteroides species isolated from 81% of anaerobic clinical infections (118). B. fragilis is not overtly invasive, but is capable of participating in intraabdominal infections in the event the mucosal wall of the intestine is disrupted. Incidences during which Bacteroides infections may be initiated include gastrointestinal surgery, perforated or gangrenous appendicitis, perforated ulcer, diverticulitis, trauma, and inflammatory bowel disease (101).
The current model for development of abdominal infections is based on the concept of synergism, during which cooperation between different species of bacteria aids in the establishment of persistent infection (121). Synergism has been most clearly established in infections involving both E. coli and B. fragilis (48), although other combinations of aerobes and anaerobes also are synergistic (125). After disruption of the intestinal wall, members of the normal flora infiltrate the normally sterile peritoneal cavity, and during the early, acute stage of infection (approximately 20 hours), the aerobes, such as E. coli, are the most active members of infection (70), establishing preliminary tissue destruction and reducing the oxidation-reduction potential of the oxygenated tissue. Once sufficient oxygen has been removed to allow the anaerobic Bacteroides to replicate, these bacteria begin to predominate during the second, chronic stage of infection (121).
The Bacteroides contribute to development of a synergistic infection in three ways: stimulation of abscess formation, reduced phagocytosis by polymorphonuclear leukocytes (PMN's), and inactivation of antibiotics by b lactamase production (121). Abscess formation is a major complication of intestinal infections, and results in the formation of a fibrous membrane surrounding a mass of cellular debris, dead PMN's, and a mixed population of bacteria. If not removed, the abscess will expand, possibly causing intestinal obstruction, erosion of resident blood vessels, and ultimately fistula formation (121). Abscesses may also metastasize, resulting in bacteremia and disseminated infection (101). Formation of the abscess is a pathological response of the immune system to the presence of the Bacteroides capsular polysaccharide. B. fragilis is the only bacterium that has been shown to induce abscess formation as the sole infecting organism (68). Purified capsule can stimulate formation of a histologically identical abscess, indicating that it is this component of the bacterium which stimulates the host immune system to deposit fibrin, forming the outer membrane of the abscess. The Bacteroides capsule has been shown to have an unusual structure, composed of repeating units of two distinct polysaccharides, each of which contains exposed positively and negatively charged side-chains (112). Most bacterial polysaccharides stimulate an antibody-mediated immune response, but the B. fragilis capsule stimulates a T cell-mediated response (69, 100, 124). Presumably, the intention of the cell-mediated immune response is to wall off the infection and protect the host from dissemination, but in fact, formation of an abscess protects the Bacteroides and neighboring bacteria from exposure to high concentrations of antibiotics and further attack from the immune system.
Another important synergistic virulence factor of B. fragilis is the ability to inhibit phagocytosis. Once the Bacteroides actively begin to replicate, they are able to interfere with attack by the immune system in two ways. First, production of the capsule itself is able to reduce the ability of the PMN's to phagocytose the bacterial cells (67, 86). Secondly, the Bacteroides are able to secrete an as yet uncharacterized factor which degrades complement proteins, and thus inhibits both chemotaxis of PMN's and opsonization of itself and neighboring bacteria (19, 45, 91).
A final contribution of the Bacteroides to a successful synergistic infection is the production of b-lactamase. Most Bacteroides strains express constitutive b-lactamase activity (66); the enzyme is extra-cellular, and thus is capable of diffusing within an abscess or other site of infection. Production of extra-cellular b-lactamases has been shown to protect other organisms in the vicinity during a mixed infection (34).
These bacteria have several other features that contribute to their pathogenicity. The Bacteroides are among the most aerotolerant of anaerobes, able to tolerate atmospheric concentrations of oxygen for up to three days (111). During initiation of an intraabdominal infection, oxygen tolerance is believed to allow the bacteria to survive in the oxygenated tissue of the abdominal cavity until E. coli and other synergistic organisms are able to reduce the redox potential at the site of infection. Additionally, this oxygen tolerance may help in surviving free radical production by the immune system PMNs. Bacteroides have been found to encode two major oxidative stress response genes, catalase (88) and superoxide dismutase (32), as well as approximately 28 other oxygen-induced proteins (87).
Although a commensal organism, Bacteroides can occasionally cause diarrhea. Strains of Bacteroides isolated from some patients with undiagnosed diarrhea were found to be enterotoxigenic, and in patients less than three years age they were associated with intestinal cramping, vomiting, and bloody stools (62). The purified toxin, fragilysin, was found to be a metalloprotease capable of hydrolysing gelatin, actin, tropomyosin, and fibrinogen (60). In a study comparing the frequency of B. fragilis enterotoxigenic and non-enterotoxigenic bacteria involved in various infection sites, the enterotoxic strains were found in higher frequencies in bacteremias (47). It is possible that fragilysin is involved in releasing the organism from an abscess or other site of infection and allowing it to enter the blood stream, thus disseminating infection throughout the body.

B. fragilis is the most common anaerobic organism isolated from clinical infections, and untreated has a mortality rate of 60% (65). This mortality rate can be greatly improved, however, with use of appropriate antimicrobial therapy (84). The Bacteroides are potentially resistant to a broad range of antibiotics, and resistance to a given antimicrobial can vary greatly between institutions. Resistance to any antimicrobial agent may occur by three mechanisms: altered target binding affinity, decreased permeability for the antibiotic, or the presence of an inactivating enzyme (80). The Bacteroides are adept at antimicrobial evasion, and may use any or all of the above mechanisms to thwart effective clinical therapy.
Antimicrobial agents may target several areas of bacterial physiology: protein translation, nucleic acid synthesis, folic acid metabolism, or cell wall synthesis (63). Protein synthesis inhibitors bind either the 30s subunit of the ribosome (aminoglycosides, tetracycline), or the 50s subunit (macrolides, lincosamides, chloramphenicol) (31). Bacteroides are inherently resistant to aminoglycosides, as uptake of this drug is energy dependent, and requires an oxygen or nitrate dependent electron transport chain which is lacking in these anaerobes (9). The Bacteroides have acquired resistances to the other protein synthesis inhibitors; resistance to clindamycin/erythromycin (macrolide-lincosamide antibiotics), and tetracycline will be discussed as pertinent examples.
Resistance to clindamycin and erythromycin has slowly but steadily increased over the last 20 years (80). In the early 1970's, all clinical isolates tested were susceptible to these antibiotics (49, 58), but late in the decade the first reports of resistance were beginning to surface (35). Three closely related genes were identified that conferred both clindamycin and erythromycin resistance in Bacteroides (33, 82, 103). These genes are similar to macrolide/lincosamide/ streptogramin resistance genes in gram positive organisms, implying that they may confer resistance by the same mechanism, namely methylation of the ribosome target site (80). Clindamycin and erythromycin resistance has been shown to be transferable between Bacteroides species, either in association with a conjugative plasmid (76, 110, 117), or a chromosomal element (54). Resistance determinants in the chromosome are often associated with tetracycline resistance, and in such instances both are cotransferred in association with a conjugative transposon (102). Resistance to to these antimicrobials in Bacteroides clinical isolates is variable from one clinical setting to another; currently resistance rates average 6%, although rates as high as 22% have been reported (107).
Tetracycline was once the first-line antibiotic for treatment of anaerobic infections. Antibiotic resistance surveys from the 1950's indicated all strains of Bacteroides were susceptible to tetracycline (30), and this antibiotic remained a frequent treatment of anaerobic infections throughout the 1960's. By the early 1970's, however, significant numbers of resistant organisms were appearing in clinical infections (49, 58), and today nearly all Bacteroides isolates are resistant, (80-90%) (80). Tetracycline resistance in the Bacteroides is attributable almost exclusively to the presence of the tetQ gene (26), which encodes a protein that is believed to alter the ribosome target site for the antibiotic (64). Bacteroides tetQ genes from several sources have been sequenced, revealing that this gene is only distantly related to other ribosomal protection genes tetM and tetO (80). The tetQ gene is chromosomally located, but can be transferred by a conjugation-like mechanism (55, 77, 78, 106). This transfer has been attributed to the presence of the tetQ gene on large, conjugative transposons called Tcr elements.
Another class of antimicrobials used to treat anaerobic infections are the Bacteroides, the most common mechanism of resistance to these compounds is production of Bacteroides species are primarily cephalosporinases, directed against the penicillin-derived cephalosporins originally developed in the 1960's for treatment of gram-negative organisms (51).
At least 90% of all Bacteroides species encode a chromosomal B. fragilis (90) and B. uniformis (104) showed them to be members of the Ambler class A
In addition to the endogenous Bacteroides also possess enzymes with activity against extended-spectrum cephalosporins (cefoxitin), and carbapenems (imipenem) (13, 23, 51). The cefoxitin resistance gene, cfxA (71), has been shown to be distantly related to the B. fragilis endogenous cepA (90). Cefoxitin was first introduced in the 1970's, and by 1980, the non-fragilis species showed significant resistance to this antimicrobial, as high as 84% in B. ovatus (35). In general, resistance to cefoxitin has remained low in B. fragilis, with rates remaining in the range of 3-6% (2, 107), although percentages as high as 11% have been reported (12).

1.Aldridge, K. E. 1995. The occurance, virulence, and antimicrobial resistance of anaerobes in polymicrobial infections. The American Journal of Surgery. 169(suppl 5A):2s-7s.
2.Aldridge, K. E., and W. D. Johnson. 1997. A comparision of susceptibility results of the Bacteroides fragilis group and other anaerobes by traditional MIC results and statistical methods. Journal of Anitmicrobial Chemotherapy. 39:319-324.
3.Ambler, R. P. 1980. The structure of B-lactamases. Philosphical Transactions of the Royal Society of London Ser. B. 289:321-331.
4.Baggio, L., and M. Morrison. 1996. The NAD(P)H-utilizing glutamate dehydrogenase of Bacteroides thetaiotaomicron belongs to enzyme family I, and its activity is affected by trans-acting gene(s) positioned downstream of gdhA. J. Bacteriol. 178:7212-7220.
5.Bokkenheuser, V. 1993. The friendly anaerobes. Clin. Infect. Dis. 16(suppl):s427-434.
6.Booth, S. J., R. L. Van Tassell, J. L. Johnson, and T. D. Wilkens. 1979. Bacteriophages of Bacteroides. Reviews of Infectious Diseases. 1:325-336.
7.Brown, J. P. 1988. Hydrolysis of glycosides and esters, p. 109-144. In I. R. Rowland (ed.), Role of the gut flora in toxicity and cancer. Academic Press, London.
8.Bry, L., P. G. Falk, T. Midtvedt, and J. I. Gordon. 1996. A model of host microbial interactions in an open mammalian ecosystem. Science. 273:1380 1383.
9.Bryan, L. E., S. K. Kowland, and H. M. Van Den Elyen. 1979. Mechanism of aminoglycoside antibiotic resistance in anaerobic bacteria: Clostridium perfringens and Bacteroides fragilis. Antimicrob. Agents Chemother. 15:7-13.
10.Cheng, Q., M. C. Yu, A. R. Reeves, and A. A. Salyers. 1995. Identification and characterization of a Bacteroides gene, csuF, which encodes an outer membrane protein that is essential for growth on chondroitin sulfate. J. Bacteriol. 177:3721-3727.
11.Croucher, S. C., A. P. Houston, C. E. Bayliss, and R. J. Turner. 1983. Bacterial populations associated with different regions of the colon wall. Appl. Environ. Microbiol. 45:1025-1033.
12.Cuchural Jr, G. J., F. P. Tally, N. V. Jacobus, K. Aldridge, T. Cleary, S. M. Finegold, G. HIll, P. Iannini, J. P. O'Keefe, C. Pierson, D. Crook, T. Russo, and D. Hecht. 1988. Susceptibility of the Bacteroides fragilis group in the United States: analysis by site of isolation. Antimicrob. Agents Chemother. 32:717-722.
13.Cuchural Jr, G. J., F. P. Tally, N. V. Jacobus, P. K. Marsh, and J. W. Mayhew. 1983. Cefoxitin inactivation by Bacteroides fragilis. Antimicrob. Agents Chemother. 32:717-722.
14.Cuchural Jr., G. J., F. P. Tally, J. R. Storey, and M. H. Malamy. 1986. Transfer of B-lactamase associated cefoxitin resistance in Bacteroides fragilis. Antimicrob. Agents Chemother. 29:918-920.
15.Cummings, J. H. 1995. Short chain fatty acids, p. 101-130. In G. R. Gibson and G. T. Macfarlane (ed.), Human colonic bacteria: role in nutrition, physiology, and pathology. CRC Press, Boca Raton.
16.D'Elia, L. N., and A. A. Salyers. 1996. Contribution of a neopullinase, a pullinase, and an alpha-glucosidase to growth of Bacteroides thetaiotaomicron on starch. J. Bacteriol. 178:7173-7179.
17.Davies, J. 1994. Inactivation of antibiotics and the dissemination of resistance genes. Science. 264:375-382.
18.Degnan, B. A., S. Macfarlane, M. E. Quigley, and G. T. Macfarlane. 1997. Starch utilization by Bacteroides ovatus isolated form the human large intestine. Curr. Microbiol. 34:290-296.
19.Dijkmans, B. A. C., P. C. Leigh, A. G. P. Braat, and R. van Furth. 1985. Effect of bacterial competition on the opsonization, phagocytosis, and intracellular killing of microorganisms by granulocytes. Infect. Immun. 49:219-224.
20.Drasar, B. S., and B. I. Duerden. 1991. Anaerobes in the normal flora of man, p. 162-179. In B. I. Duerden and B. S. Drasar (ed.), Anaerobes in human disease. Wiley-Liss, New York.
21.Edenharder, R. 1984. Dehydroxylation of cholic acid at C12and epimerization at C5 and C7 by Bacteroides species. J. Ster. Chem. 21:413-420.
22.Edwards, R., and D. Greenwood. 1992. An investigation of B-lactamases from clinical isolates of Bacteroides species. J. Med. Microbiol. 36:89-95.
23.Eley, A., and D. Greenwood. 1986. Beta-lactamases of type culture strains of the Bacteroides fragilis group and of strains that hydrolyse cefoxitin and imipenem. J. Med. Microbiol. 21:49-57.
24.Eyssen, H., and P. H. Caenepeel. 1988. Metabolism of fats, bile acids, and steroids, p. 263-286. In I. R. Rowland (ed.), role of the gut flora in toxicity and cancer. Academic Press, London.
25.Finegold, S. M., V. L. Sutter, and G. E. Mathisen. 1983. Normal indigenous intestinal flora, p. 3-31. In D. J. Hentges (ed.), Human intestinal microflora in health and disease. Academic Press, New York.
26.Fletcher, H. M., and M. H. Malamy. 1991. Molecular survey of clindamycin and tetracycline resistance determinants in Bacteroides species. Antimicrob. Agents Chemother. 35:2415-2418.
27.Gherna, R., and C. R. Woese. 1992. A partial phylogenetic analysis of the "Flavobacter-Bacteroides" phylum: basis for taxonomic restructuring. Syst. Appl. Microbiol. 15:513-521.
28.Gibson, S. A., and G. T. Macfarlane. 1988. Characterization of proteases formed by Bacteroides fragilis. J. Gen. Microbiol. 134:2231-2240.
29.Gibson, S. A., and G. T. Macfarlane. 1988. Studies on the proteolytic activity of Bacteroides fragilis. J. Gen. Microbiol. 134:19-27.
30.Gillespie, W. A., and J. Guy. 1956. Bacteroides in intra-abdominal sepsis. Lancet. 1:1039-1041.
31.Greenwood, D., B. Watt, and B. I. Duerden. 1991. Antibiotics and anaerobes, p. 415-429. In B. I. Duerden and B. S. Drasar (ed.), Anaerobes in Human Disease. Wiley-Liss, New York.
32.Gregory, E. M., and C. H. Dapper. 1983. Isolation of iron-containing super oxide dismutase from Bacteroides fragilis: reconstitution as a Mn-containing enzyme. Arch. Biochem. Biophys. 220:293-300.
33.Halula, M. C., S. Manning, and F. L. Macrina. 1991. Nucleotides sequence of ermFU, a macrolide-lincosamide-streptogramin (MLS) resistance gene encoding an RNA methlyase from the conjugal element of Bacteroides fragilis V503. Nucleic Acids Res. 19:3453.
34.Hamilton-Miller, J. M. T. 1981. Indirect pathogenicity. J. Antimicrob. Chemother. 7:307-308.
35.Hansen, S. L. 1980. Variation in susceptibility patterns within the Bacteroides fragilis group. Antimicrob. Agents Chemother. 17:686-690.
36.Hentges, D. J. 1989. Anaerobes as normal flora, p. 37-53. In S. M. Finegold and W. L. George (ed.), Anaerobic infections in humans. Academic Press, San Diego.
37.Hentges, D. J. 1983. Role of the intestinal flora in host defense against infection, p. 311-331. In D. J. Hentges (ed.), Human intestinal microflora in health and disease. Academic Press, New York.
38.Hespell, R. B., and C. J. Smith. 1983. Utilization of nitrogen sources by gastrointestinal tract bacteria, p. 167-187. In D. J. Hentges (ed.), Human intestinal microflora in health and disease. Academic Press, New York.
39.Holdeman, L. V. R., R. W. Kelly, and W. E. C. Moore. 1984. Genus I. Bacteroides Castellani and Chalmers 1919. 959AL, p. 604-631. In N. R. Krieg and J. G. Holt (ed.), Bergey's manual of systematic bacteriology, vol. 1. The Williams and Wilkins Co., Baltimore.
40.Huhtanen, C. M. 1979. Bile acid inhibition of Clostridium botulinum. Appl. Environ. Microbiol. 38:216-218.
41.Hylemon, P. B., and T. L. Glass. 1983. Biotransformation of bile acids and cholesterol by the intestinal microflora, p. 189-213. In D. J. Hentges (ed.), Human intestinal microflora in health and disease. Academic Press, New York.
42.Hylemon, P. B., and P. V. Jr. Phibbs. 1974. Evidence against the presence of cyclic AMP and related enzymes in selected strains of Bacteroides fragilis. Biochem. Biophys. Res. Commun. 60:88-95.
43.Hylemon, P. B., and J. A. Sherrod. 1975. Multiple forms of 7ð hydroxysteroid dehydrogenases in selected strains of Bacteroides fragilis. J. Bacteriol. 122:418-424.
44.Hylemon, P. B., J. L. Young, R. F. Roadcap, and P. V. J. Phibbs. 1977. Uptake and incorporation of glucose and mannose by whole cells of Bacteroides thetaiotaomicron. Appl. Environ. Microbiol. 34:488-494.
45.Ingham, H. R., P. R. Sisson, D. Tharagonnet, J. B. Selkon, and A. A. Codd. 1977. Inhibition of phagocytosis in vitro by obligate anaerobes. Lancet. ii:1252 1254.
46.Johnson, J. L. 1978. Taxonomy of the Bacteroides.I. Deoxyribonucleic acid homologies among Bacteroides fragilis and other saccharolytic Bacteroides species. Int. J. Syst. Bacteriol. 28:245-256.
47.Kato, N., H. Kato, K. Wantanabe, and K. Ueno. 1996. Association of enterotoxigenic Bacteroides fragilis with bacteremia. Clinical Infectious Disease. 23(supple 1):s83-86.
48.Kelly, M. J. 1978. The quantitative and histological demonstration of pathogenic synergy between Escherichia coli and Bacteroides fragilis in guinea-pig wounds. J. Med. Microbiol. 11:513-523.
49.Kislak, J. W. 1972. The susceptiblity of Bacteroides fragilis to 24 antibiotics. Journal of Infectious Diseases. 125:295-298.
50.Lambe, D. W., Jr. 1974. Determination of Bacteroides melaninogenicus serogroups by fluorescent antibody staining. Appl. Microbiol. 28:561-567.
51.Livermore, D. M. 1993. Carbapenemases: the next generation of B lactamases. ASM News. 59:129-135.
52.Macfarlane, G. T., and S. Macfarlane. 1997. Human colonic microbiota: ecology, physiology and metabolic potential of intestinal bacteria. :3-9.
53.Maier, B. R., and D. J. Hentges. 1972. Experimental Shigella infections in laboratory animals.I. Antagonism by normal flora components in gnotobiotic mice. Infect. Immun. 6:168-173.
54.Malamy, M. H., and F. P. Tally. 1981. Mechanisms of drug-resistance transfer in Bacteroides fragilis. J. Antimicrob. Chemother. 8(suppl D):59-75.
55.Mays, T. D., C. J. Smith, R. A. Welch, C. Delfini, and F. L. Macrina. 1982. Novel antibiotic resistance transfer in Bacteroides. Antimicrob. Agents Chemother. 21:110-118.
56.McNeil, N. I., J. H. Cummings, and W. P. T. James. 1978. Short chain fatty acid absorbtion by the human large intestine. Gut. 19:819-822.
57.Meynell, G. G. 1963. Antibacterial mechanisms of the mouse gut.II. The role of Eh and volatile fatty acids in the normal gut. Brit. J. Exp. Path. 44:209 219.
58.Mitchell, A. A. B. 1973. Incidence and isolation of Bacteroides species from clinical material and their sensitivity to antibiotics. J. Clin. Path. 26:738-741.
59.Miyagawa, E., R. Azuma, and T. Suto. 1979. Distribution of sphingolipids in Bacteroides species. J. Gen. Appl. Microbiol. 25:41-52.
60.Moncrief, J. S., R. Obiso Jr. , L. A. Banoso, J. J. Kling, R. L. Wright, R. L. Van Tassell, D. M. Lyerly, and T. D. Wilkins. 1995. The enterotoxin of Bacteroides fragilis is a metalloprotease. Infect. Immun. 63:175-181.
61.Moore, W. E. C., and L. V. Holdeman. 1974. Human fecal flora: the normal flora of 20 Japenese-Hawaiians. Appl. Microbiol. 27:961-79.
62.Myers, L. L., D. S. Shoop, L. L. Stackhouse, F. S. Newman, R. J. Flaherty, G. W. Letson, and R. B. Sack. 1987. Isolation of enterotoxigenic Bacteroides fragilis from humans with diarrhea. J. Clin. Microbiol. 25:2330-2333.
63.Neu, H. C. 1995. Emergance and mechanisms of bacterial resistance in surgical infections. The American Journal of Surgery. 169(suppl 5A):13s-20s.
64.Nikolich, M. P., N. B. Shoemaker, and A. A. Salyers. 1992. A Bacteroides tetracycline resistance gene represents a new class of ribosome protection tetracycline resistance. Antimicrob. Agents Chemother. 36:1005-1012.
65.Nobles Jr., E. R. 1973. Bacteroides infections. Anal. Surg. 177:601-606.
66.Nord, C. E., and D. Olsson-Liljequist. 1981. Resistance to B-lactam antibiotics in Bacteroides species. J. Antimicrob. Chemother. 8(suppl):33-42.
67.Okuda, K., and I. Takozoe. 1973. Antiphagocytic effects of the capsular structure of a pathogenic strain of Bacteroides melaninogenicus. Bull. Tokyo Dent. Col. 14:99-104.
68.Onderdonk, A. B., D. L. Kasper, R. L. Cisneros, and J. G. Bartlett. 1977. The capsular polysaccharide of Bacteroides fragilis as a virulence factor: Comparision of the pathogenic potential of encapsulated and unencapsulated strains. J. Infect. Dis. 136:82-89.
69.Onderdonk, A. B., R. B. Markham, D. F. Zaleznik, R. L. Cisneros, and D. L. Kasper. 1982. Evidence for T cell-dependent
immunity to Bacteroides fragilis in an intraabdominal abscess model. J. Clin. Invest. 69:9-16.
70.Onderdonk, A. B., W. M. Weinstein, N. M. Sullivan, J. G. Bartlett, and S. L. Gorbach. 1974. Experimental intra-abdominal abscess in rats: quantitative bacteriology of infected animals. Infect. Immun. 10:1256-1258.
71.Parker, A. C., and C. J. Smith. 1993. Genetic and biochemical analysis of a novel Ambler class A beta-lactamase responsible for cefoxitin resistance in Bacteroides species. Antimicrob. Agents Chemother. 37:1028-1036.
72.Paster, B. J., F. E. Dewhirst, I. Olsen, and G. J. Fraser. 1994. Phylogeny of Bacteroides, Prevotella, and Porphyromonas spp. and related bacteria. J. Bacteriol. 176:725-732.
73.Paster, B. J., W. Ludwig, W. G. Weisburg, E. Stackebrandt, R. B. Hespell, C. M. Hahn, H. Reichenbach, K. O. Stetter, and C. R. Woese. 1985. A phylogenetic grouping of the Bacteroides, Cytophagas, and certain Flavobacteria. Syst. Appl. Microbiol. 6:34-42.
74.Piddock, L. V. J., and R. Wise. 1987. Cefoxitin resistance in Bacteroides species: evidence indicating two mechanisms causing decreased susceptibility. J. Antimicrob. Chemother. 19:161-170.
75.Prins, R. A. 1977. Biochemical activities of gut microorganisms, p. 73-183. In R. T. J. Clarke and T. Bauchop (ed.), Microbial ecology of the gut. Academic Press, New York.
76.Privetera, G., A. Dublanchet, and M. Sebald. 1979. Transfer of multiple antibiotic resistance between subspecies of Bacteroides fragilis. Journal of Infectious Diseases. 139:97-101.
77.Privitera, G., M. Sebald, and F. Fayolle. 1979. Common regulatory mechanism of expression and conjugative ability of a tetracycline resistance plasmid in Bacteroides fragilis. Nature. 278:657-659.
78.Rashtchian, A., G. R. Dubes, and S. J. Booth. 1982. Tetracycline-inducible transfer of tetracycline resistance in Bacteroides fragilis in the absence of detectable plasmid DNA. J. Bacteriol. 150:141-147.
79.Rashtchian, A., G. R. Dubes, and S. J. Booth. 1982. Transferable resistance to cefoxitin in Bacteroides thetaiotaomicron. Antimicrob. Agents Chemother. 22:701-703.
80.Rasmussen, B. A., K. Bush, and F. P. Tally. 1997. Antimicrobial resistance in anaerobes. Clin. Infect. Dis. 24(suppl 1):s110-120.
81.Rasmussen, B. A., K. Bush, and F. P. tally. 1993. Antimicrobial resistance in Bacteroides. Clin. Infect. Dis. 16(suppl 4):s390-s400.
82.Rasmussen, J. L., D. A. Odelson, and F. L. Macrina. 1986. Complete nucleotide sequence and transcription of ermF, a macrolide-lincosamide streptogramin B resistance determinant from Bacteroides fragilis. J. Bacteriol. 168:523-533.
83.Reddy, N. R., J. K. Palmer, M. D. Pierson, and R. J. Bothast. 1984. Intracellular glycosidases of human colon Bacteroides ovatus B4-11. Appl. Environ. Microbiol. 48:890-892.
84.Redondo, M. C., M. D. J. Arbo, J. Grindlinger, and D. R. Snydman. 1995. Attributable mortality of bacteremia associted with the Bacteroides fragilis group. Clin. Infect. Dis. 20:1492-1496.
85.Reeves, A. R., G. R. Wang, and A. A. Salyers. 1997. Characterization of four outer membrane proteins that play a role in utilization of starch by Bacteroides thetaiotaomicroon. J. Bacteriol. 179:643-649.
86.Reid, J. H., and S. Patrick. 1984. Phagocytic and serum killing of capsulate and non-capsulate Bacteroides fragilis. J. Med. Microbiol. 17:247-257.
87.Rocha, E. R., T. Selby, J. P. Coleman, and C. J. Smith. 1996. Oxidative stress response in an anaerobe, Bacteroides fragilis: a role for catalase protection against hydrogen peroxide. J. Bacteriol. 178:6895-6903.
88.Rocha, E. R., and C. J. Smith. 1995. Biochemical and genetic analysis of a catalase from the anaerobic bacterium Bacteroides fragilis. J. Bacteriol. 177:3111-3119.
89.Rogers, M. B., T. K. Bennett, C. M. Payne, and C. J. Smith. 1994. Insertional inactivation of cepA leads to high-level B-lactamase expression in Bacteroides fragilis clinical isolates. J. Bacteriol. 176:4376-4384.
90.Rogers, M. B., A. C. Parker, and C. J. Smith. 1993. Cloning and characterization of the endogenous cephalosporinase gene, cepA, from Bacteroides fragilis reveals a new subgroup of Ambler class A B-lactamases. Antimicrob. Agents Chemother. 37:2391-2400.
91.Rotstein, O., T. L. Pruett, J. J. Sorenson, V. D. Fiegal, R. D. Nelson, and R. L. Simmons. 1986. A Bacteroides by-product inhibits human polymorhonuclear leukocyte function. Arch. Surg. 121:81-88.
92.Salyers, A. A., and J. A. Z. Leedle. 1983. Carbohydrate metabolism in the human colon, p. 129-146. In D. J. Hentges (ed.), Human intestinal microflora in health and disease. Academic Press, New York.
93.Salyers, A. A., M. O'Brian, and S. F. Kotarski. 1982. Utilzation of chondroitin sulfate by Bacteroides thetaiotaomicron growing in carbohydrate limited continuous culture. J. Bacteriol. 150:1008-1015.
94.Salyers, A. A., J. R. Vercellotti, S. E. H. West, and T. D. Wilkins. 1977. Fermentation of mucin and plant polysaccharides by strains of Bacteroides from the human colon. Appl. Environ. Microbiol. 33:319-322.
95.Scholle, R. R., H. E. Steffen, H. J. K. Goodman, and D. R. Woods. 1990. Expression and regulation of a Bacteroides fragilis sucrose utilization system cloned in Escherichia coli. Appl. Environ. Microbiol. 56:1944-1948.
96.Shah, H. N. 1992. The genus Bacteroides and related taxa, p. 3593-3607. In A. Balows, H. G. Truper, M. Dworkin, W. Harder, and K.-H. Schleifer (ed.), The Prokaryotes, 2nd ed, vol. 4. Springer-Verlag, New York.
97.Shah, H. N., and M. D. Collins. 1990. Prevotella, a new genus to include Bacteroides melaninogenicus and related species formerly classified in the genus Bacteroides. Int. J. Syst. Bacteriol. 40:205-208.
98.Shah, H. N., and M. D. Collins. 1988. Proposal for reclassification of Bacteroides asaccharolyticus, Bacteroides gingivalis, and Bacteroides endodontalis in a new genus, Porphyromonas. Int. J. Syst. Bacteriol. 38:128 131.
99.Shah, H. N., and M. D. Collins. 1989. Proposal to restrict the genus Bacteroides (Castellani and Chalmers) to Bacteroides fragilis and closely related species. Int. J. Syst. Bacteriol. 39:85-87.
100.Shapiro, M. E., D. L. Kasper, D. F. Zaleznik, S. Spriggs, A. B. Onderdonk, and R. W. Finberg. 1986. Cellular control of abscess formation: Role of T cells in the regulation of abscesses formed in response to Bacteroides fragilis. J. Immun. 137:314-346.
101.Sheehan, G., and G. Harding. 1989. Intraperitoneal infections, p. 349-384. In S. M. Finegold and W. L. George (ed.), Anaerobic infections in humans. Academic Press, San Diego.
102.Shoemaker, N. B., R. D. Barber, and A. A. Salyers. 1989. Cloning and characterization of a Bacteroides conjugal tetracycline-erythromycin resistance element by using a shuttle cosmid vector. J. Bacteriol. 171:1294-1302.
103.Smith, C. J. 1987. Nucleotide sequence analysis of Tn4551: use of ermFS operon fusions to detect promoter activity in Bacteroides fragilis. J. Bacteriol. 169:4589-4596.
104.Smith, C. J., T. K. Bennett, and A. C. Parker. 1994. Molecular and genetic analysis of the Bacteroides uniformis cephalosporinase gene, cblA, encoding the species-specific B-lactamase. Antimicrob. Agents Chemother. 38:1711-1715.
105.Smith, C. J., and A. C. Parker. 1993. Identification of a circular intermediate in the transfer and transposition of Tn4555, a mobilizable transposon from Bacteroides spp. J. Bacteriol. 175:2682-2691.
106.Smith, C. J., R. A. Welch, and F. L. Macrina. 1982. Two independent conjugal transfer systems operating in Bacteroides fragilis V479-1. J. Bacteriol. 151:281-287.
107.Snydman, D. R., L. McDermott, G. J. Cuchural, D. W. Hecht, P. B. Iannini, L. J. Harrell, S. G. Jenkins, J. P. O'Keefe, C. L. Pierson, J. D. Rihs, V. L. Yu, S. M. Finegold, and S. L. Gorbach. 1997. Analysis of trends in antimicrobial resistance patterns among clinical isolates of Bacteroides fragilis group species from 1990 to 1994. Clin. Infect. Dis. 23(suppl 1):s54-65.
108.Southern, J. A., J. R. Parker, and D. R. Woods. 1986. Expression and purification of glutamine synthetase cloned from Bacteroides fragilis. J. Gen. Microbiol. 132:2827-2835.
109.Stellwag, E. J., and P. B. Hylemon. 1976. Purification and characterization of bile salt hydrolase from Bacteroides fragilis subsp fragilis. Biochemica Biophysica Acta. 452:165-176.
110.Tally, F. P., D. R. Snydman, S. L. Gorbach, and M. H. Malamy. 1979. Plasmid-mediated, transferable resistance to clindamycin and erythromycin in Bacteroides fragilis. Journal of Infectious Diseases. 139:83-88.
111.Tally, F. P., P. R. Stewart, V. L. Sutter, and J. E. Rosenblatt. 1975. Oxygen tolerance of fresh clinical isolates. J. Clin. Microbiol. 1:161-164.
112.Tzianabos, A. O., A. B. Onderdonk, B. Rosner, R. L. Cisneros, and D. L. Kasper. 1993. Structural features of polysaccharides that induce intra abdominal abscesses. Science. 262:416-419.
113.Van den Eynde, H., R. De Baere, H. N. Shah, S. E. Gharbia, G. E. Fox, J. Michalik, Y. Van de Peer, and R. De Wachter. 1989. 5S ribosomal ribonucleic acid sequences in Bacteroides and Fusobacteria: Evolutionary relationships within these genera and among Eubacteria in general. International Journal of Systemic Bacteriology. 39:78-84.
114.van der Waaij, D., J. M. Berghuis de Vries, and J. E. C. Lekkerkerk van der Wees. 1971. Colonization resistance of the digestive tract in conventional and antibiotic-treated mice. J. Hyg. 69:405-411.
115.Veillon, M. H., and H. Zuber. 1898. Recherches sur quelques microbes strictement anaerobes et leur cole en pathologie. Arch. Exp. Med. Path. Anat. 10:517-545.
116.Weisburg, W. G., Y. Oyaizu, H. Oyaizu, and C. R. Woese. 1985. Natural relationship between Bacteroides and Flavobacteria. J. Bacteriol. 164:230-236.
117.Welch, R. A., K. R. Jones, and F. L. Macrina. 1979. Transferable lincosamide-macrolides resistance in Bacteroides. Plasmid. 2:261-268.
118.Werner, H. 1974. Differentiation and medical importance of saccharolytic intestinal anaerobes. Arz. Forsch. 24:340-343.
119.Wexler, H. M., and S. Halebian. 1990. Alterations to the penecillin-binding proteins on the Bacteroides fragilis group: a mechanism for non-B-lactamase mediated cefoxitin resistance. J. Antimicrob. Chemother. 26:7-20.
120.Whitehead, T. R. 1995. Nucleotides sequences of xylan-inducable xylanase and xylosidase/arabinosidase genes from Bacteroides ovatus V975. Biochemica st Biophysica Acta. 1244:239-241.
121.Willis, A. T. 1991. Abdominal sepsis, p. 197-223. In B. I. Duerden and B. S. Drasar (ed.), Anaerobes in Human Disease. Wiley-Liss, New York.
122.Wrong, O. M. 1988. Bacterial metabolism of protein and endogenous nitrogen compounds, p. 227-262. In I. R. Rowland (ed.), Role of the gut flora in toxicity and cancer. Academic Press, London.
123.Yotsuji, A., S. Minami, M. Inoue, and S. Mitsukashi. 1983. Properties of a novel B-lactamase produced by Bacteroides fragilis. Antimicrob. Agents Chemother. 24:925-929.
124.Zaleznik, D. F., R. W. Finberg, M. E. Shapiro, A. B. Onderdonk, and D. L. Kasper. 1985. A soluble suppressor T cell factor protects against experimental intraabdominal abscesses. J. Clin. Invest. 75:1023-1027.
125.Zaleznik, D. F., and D. L. KAsper. 1989. Role of bacterial virulence factors in pathogenesis of anaerobic infection, p. 81-95. In S. M. Finegold and W. L. George (ed.), Anaerobic infections in humans. Academic Press, San Diego.