Department of Microbiology, University of Leeds and The General Infirmary, Leeds LS2 9JT, UK
Received 5 November 2002; returned 20 January 2003; revised 21 March 2003; accepted 2 April 2003
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Abstract |
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Keywords: Clostridium difficile, cefotaxime, desacetylcefotaxime, gut model, gut microflora
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Introduction |
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Materials and methods |
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MacFarlane et al.26 validated a three-stage continuous culture system to investigate the effect of retention time on the ecology and metabolism of bacteria in the human colon. The system was designed to reproduce the spatial, temporal, nutritional and physicochemical characteristics of the proximal and distal bowel. The model consists of three vessels (V1, V2 and V3), operating in a weir cascade system, running into a waste unit. The model was temperature controlled (37°C), and vessels were automatically controlled at pH 5.5, 6.2 and 6.8, respectively, to reflect increasing alkalinity of the human intestine from proximal to distal. V1 had an operating volume of 280 mL, whereas V2 and V3 operated at 300 mL. Thus V1 had a high availability of substrate, allowing rapid bacterial growth, and was operated at an acidic pH, similar to events in the proximal colon. In contrast, the final vessel resembled the neutral pH, slow bacterial growth rate and low substrate availability, which is characteristic of more distal regions. V1 was supplied with growth medium, at a rate controlled by a peristaltic pump. All three vessels and the growth medium reservoir were continuously stirred and sparged with oxygen-free nitrogen. The model was allowed initially to equilibrate for 2 weeks in respect of bacterial populations, and then operated at a total retention time of 67 h (V1 = 16.7 h; V2 and V3 = 25 h).
Sample collection and inoculation of the model
Fresh faeces were collected from five healthy elderly volunteers (aged >65 years) with no history of antibiotic therapy for at least 4 weeks prior to the study. Faecal specimens were transported to the laboratory in anaerobic bags at 4°C (Becton Dickinson), whereupon they were investigated for 48 h at 37°C for the presence of C. difficile by anaerobic culture on CCEY agar, supplemented with 2% lysed horse blood and 5 mg/L lysozyme. C. difficile-negative faecal specimens were pooled to give 50 g of faeces. Pooled faeces were emulsified in pre-reduced PBS (pH 7.4) to give a 10% faecal slurry. A smooth slurry (500 mL) was obtained by stomaching (BoroLabs, Aldermaston, UK), followed by coarse filtration through sterile muslin, under anaerobic conditions. This slurry was then used to inoculate V1, V2 and V3 to two-thirds volume. Growth medium was added to V1 to give a final volume of 280 mL, whereupon the medium pump was started to allow V2 and V3 to be fed from V1.
Growth medium
Growth medium was prepared in 5 L volumes and sparged with nitrogen prior to adding to the model. The medium consisted of (g/L): peptone water 2.0, yeast extract 2.0, NaCl 0.1, K2HPO4 0.04, KH2PO4 0.04, MgSO4.7H2O 0.01, CaCl2.2H2O 0.01, NaHCO3 2.0, haemin 0.005, cysteine HCl 0.5, bile salts 0.5, arabinogalactan 1, pectin 2, starch 3. Liquid additions were made as follows: vitamin K 10 µL/L and Tween 80 0.2%. After autoclaving, glucose (0.4 g/L) and starch (3 g/L) solutions were added to the medium through a sterile filtration device. Resazurin anaerobic indicator was also added at 0.005 g/L.
Enumeration of C. difficile spores
Two millilitre samples were removed from each vessel, and 0.5 mL of each sample was treated with 0.5 mL 96% ethanol and incubated at room temperature for 1 h. Each alcohol-shocked sample was then serially diluted to 107 in pre-reduced peptone water. Twenty microlitres of each dilution was plated onto Braziers CCEY agar base, supplemented with 2% lysed blood and 5 mg/L lysozyme (no antibiotics or egg yolk were included). Plates were incubated under anaerobic conditions (10% CO2, 10% H2, 80% N2) at 37°C (Don Whitley Scientific) for 48 h. Single colonies were then counted.
Enumeration of faecal bacteria
From each 2 mL sample, 0.5 mL was serially diluted to 109 in an anaerobic cabinet in 4.5 mL pre-reduced peptone water. Twenty microlitres of each dilution was then used to inoculate the following selective agars (all agars supplied by Oxoid, Basingstoke, UK unless otherwise stated and all supplements supplied by Sigma-Aldrich unless otherwise stated): nutrient agar (total aerobes); MacConkey agar, no. 3 (lactose fermentors); WilkinsChalgren agar (total anaerobes); azide agar (Gram-positive cocci); reinforced clostridial agar supplemented with 8 mg/L novobiocin, 8 mg/L colistin (total clostridial counts); Rogosa agar (lactobacilli); Beerens agar42.5 g/L Columbia agar; 5 g/L agar technical; 5 g/L glucose; 0.5 g/L cysteine HCl (bifidobacteria); Brucella agar supplemented with 75 mg/L kanamycin; 5 mg/L haemin; 10 µL/L vitamin K1; 7.5 mg/L vancomycin; 5% laked horse blood (E and O Laboratories, Scotland). All agars were pre-reduced for 24 h prior to inoculation in an anaerobic cabinet. Agar plates were incubated for 48 h in an anaerobic cabinet. After incubation, colonies were counted and identified to genus level on the basis of colony morphology, Gram reaction and microscopic appearance.
C. difficile cytotoxin quantification
Five hundred microlitres of each 2 mL sample was centrifuged at 16 000g and the supernatant removed. The supernatant was then serially diluted in PBS (pH 7.4) to 107. Twenty microlitres of each dilution was added to VERO cell culture monolayers prepared in 96-well microtitre trays. Cell culture toxin assays were incubated at 37°C in air with 5% CO2, and examined at 24 and 48 h under an inverted microscope. A positive reaction was indicated by cell rounding. Specific action of C. difficile cytotoxin was confirmed by parallel neutralization with Clostridium sordellii antitoxin (ProLab Diagnostics). Toxin titres were expressed as relative units (RU).
Preparation of C. difficile spores
Ten fresh blood agar plates were inoculated with the C. difficile UK epidemic strain (CDUKES), PCR ribotype I,29 and incubated in an anaerobic cabinet for 7 days at 37°C. All growth was removed from all the plates with a sterile swab, and resuspended in 1 mL of sterile water. An equal volume of 95% ethanol was mixed with the suspension, which was then incubated at room temperature for 1 h. Suspensions were then centrifuged at 3000g and resuspended in sterile water. Viable C. difficile spores were enumerated by culturing 20 µL serial dilutions in pre-reduced PBS on Braziers CCEY lysozyme agar. This was carried out in an anaerobic environment for 48 h at 37°C. Spore suspensions were adjusted to achieve a concentration of 107 cfu/mL.
Preparation of antibiotic and metabolite
CTX and dCTX were prepared to achieve final concentrations of 20 and 40 mg/L, respectively, when added to V1. These concentrations are approximately equivalent to those found in human bile following a single 2000 mg dose of CTX.21 Both CTX and dCTX were supplied by Aventis Pharma.
Inoculation and antibiotic instillation to the gut model
After inoculation with faecal emulsion, the model was allowed to achieve steady state for 2 weeks. Bacterial populations were continuously monitored during this time. At steady state, the model was inoculated with 107 cfu/mL C. difficile spores in a 1 mL volume (period A). After 7 days, the model was inoculated with a further 107 cfu/mL C. difficile spores (in a 1 mL volume). At this point, the model was dosed at 12 h intervals with CTX 20 mg/L for 7 days (period B), followed by 7 days without CTX instillation (period C). The model was then inoculated with a further 107 cfu/mL C. difficile spores (in a 1 mL volume) and dosed at 12 h intervals with CTX 20 mg/L and dCTX 40 mg/L for 7 days (period D), followed by 7 days without CTX/dCTX instillation (period E). Bacterial populations, C. difficile spores and cytotoxin were enumerated, as described above, throughout this period. Vessels were sampled at approximately the same time each day, and always exactly at midpoint between antibiotic doses (i.e. 6 h post- and pre-dose).
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Results |
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The addition of C. difficile spores to the model did not affect viable counts of faecal bacteria in any of the three vessels (period A). Relative numbers of C. difficile spores and vegetative cells remained similar in the 7 days following addition of spores to the model. C. difficile cytotoxin was undetectable (Table 1 and Figure 1ac).
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The seven twice daily instillations of CTX to the model produced had no observable effects upon total aerobic counts and numbers of Gram-positive cocci. Slight decreases in numbers over the initial period of antibiotic instillation were observed for lactose fermenters, clostridia, bifidobacteria, Bacteroides spp. and total anaerobes, in all three vessels. Counts recovered towards the later stages of instillation to at least their original levels. Lactobacilli counts rose sharply during CTX instillation in all three vessels (period B).
C. difficile total viable cells (TVC) and spores remained at their original levels in all three vessels until the fourth day of CTX instillation (day 28), when an increase in TVC and a decrease in spores was observed in V1. This was subsequently observed in V2 on the fifth day of CTX treatment (day 29) and in V3 on the sixth day of CTX instillation (day 30). Cytotoxin titres showed no increase in V1 until day 7 of CTX instillation (day 31), whereupon a slight rise to a titre of 1 RU was observed. In V2 and V3, cytotoxin titres rose sharply on day 5 of CTX instillation (day 29), from undetectable to a titre of 4 RU, attaining their highest titres of 5 RU on day 6 of CTX exposure (day 30). The increase in TVC and decrease in spores continued until cessation of CTX instillation on day 32 (period D). At this point, TVC decreased sharply, whereas spore counts plateaued. Cytotoxin titres in V2 and V3 decreased sharply (Figure 1ac).
Following the cessation of CTX instillation, and concurrent with the fall in C. difficile numbers and cytotoxin titres, lactobacilli and bifidobacterial populations increased beyond their original numbers, whereas others (Bacteroides spp., clostridia, GPC, aerobes) maintained the levels seen during the later stages of CTX instillation (period C).
Effect of CTX and dCTX addition to the gut model (period D)
The effects upon gut bacteria of seven twice daily instillations of CTX and dCTX to the model were similar to the addition of CTX alone. A decrease in numbers of bifidobacteria in V1 in the initial period of CTX/dCTX instillation was followed 3 days later by a similar decrease in numbers of bifidobacteria in V2 and V3. Lactobacilli decreased in numbers in all three vessels during the initial stages of CXT/dCTX instillation, but recovered in the later stages. Changes in other bacterial populations were similar to those observed during instillation with CTX alone. Pre-antibiotic exposure numbers of faecal bacteria were re-established by the end of the experiment.
A profound decrease in numbers of Bacteroides spp. was observed in all three vessels (Figure 2a, period D). In V1, numbers decreased from 107 cfu/mL on the first day of CTX/dCTX instillation to undetectable levels by the fifth day of instillation (day 44). Similarly, in V2 and V3, numbers decreased from 108 and 107 cfu/mL, respectively, on the first day of CTX/dCTX instillation, to 105 and 104 cfu/mL by the last day of instillation (day 47). However, counts had re-established themselves to their original pre-exposure levels less than 48 h following cessation of therapy (Figure 2a, periods D/E). A slight decrease in numbers of bifidobacteria was also observed during antibiotic instillations, which also recovered upon cessation of antibiotic administration (Figure 2b).
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Discussion |
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Following the introduction of C. difficile spores to the model, there was no evidence of spore germination, exponential growth or toxin production prior to the introduction of CTX. The organism appeared to remain in the same quiescent state as when introduced to the model. We believe this is analogous to colonization in vivo. Since the infective dose of C. difficile spores is not known, this study used a relatively heavy spore inoculum in order to ensure colonization. The fact that such a high concentration of spores was used and yet no active germination, growth or toxin production was seen is evidence in favour of this period representing colonization, as opposed to active disease.
The possible influence of antibiotic metabolites upon C. difficile and gut flora has largely been overlooked. A study from the USA compared the effects of clindamycin and its metabolites in the hamster model of CDI, finding no correlation between antibacterial potency and AACD50 (toxin lethal to 50% of animals).32 However, this study was not controlled, either in terms of infective dose or infecting strain of C. difficile. Our study is the first to examine the effects of cefotaxime and its metabolites on C. difficile and gut flora. Previous investigations in test-tube models have failed to explain the particular propensity of cefotaxime towards C. difficile infection, but have also failed to account for either the level of antibiotic penetration into faeces or the possible presence of metabolites. The model was exposed to concentrations of CTX and dCTX similar to the peak levels found in bile, where dCTX is found in excess of CTX in an 2:1 ratio, and also aims to mimic a 7 day 2 g twice-daily CTX course of therapy. Although CTX and dCTX concentrations were not determined in each vessel, the addition of the drug(s) at peak level, and at regular dosing intervals, was designed to reflect the fluctuation of antibiotic concentrations within the gut. In this way we hoped to reflect the high levels achieved in the gut lumen shortly after administration, followed by a slow decline in levels as the contents of V1 fed into V2, and subsequently V3, became more dilute.
Treatment with CTX alone produced a slight decrease in numbers of most bacterial groups (excepting C. difficile), followed rapidly by re-establishment of original numbers. Treatment with CTX/dCTX produced reduced numbers of bifidobacteria in V2 and V3, and had a profoundly deleterious effect upon Bacteroides spp. in V1, with a less marked effect in V2 and V3. This is in agreement with the recent report investigating the bacterial populations in faecal samples from healthy young adults, elderly subjects and elderly patients with C. difficile-associated diarrhoea (CDAD).33 Patients with CDAD had a greater diversity of facultative species in gut flora, such as lactobacilli and clostridia, but greatly reduced numbers of Bacteroides spp., Prevotella spp. and bifidobacteria.33,34 In addition, comparative examinations of faecal bacterial populations have shown a decrease in bifidobacterial and Bacteroides spp. diversity with age. In particular, only one Bacteroides spp. was isolated from this group. There were no data available, however, on predisposing antibiotic exposure of these patients. An early study by Ambrose et al.16 noted that antibiotics associated with the emergence of C. difficile (latamoxef, cefotaxime, cefoxitin, ceftriaxone and cefotetan) were also particularly active against B. fragilis.
The potential role of bifidobacteria in colonization resistance has been examined.3538 Ouwehand et al.35 found that four bifidobacterial strains adhered significantly less to mucus from elderly individuals compared with the other subjects tested. Some studies have reported inhibitory effects of bifidobacteria directed against Escherichia coli, Clostridium perfringens, lecithinase-producing clostridia and Bacteroides spp.,3638 and Bifidobacterium longum culture supernatant was recently shown to neutralize the effects of E. coli vero toxin.39 It is possible that a similar effect may be achieved against C. difficile cytotoxins. Bifidobacterium bifidum was shown to produce biotin extracellularly when cultured on oligosaccharide-containing media. Biotin-limited conditions were associated with increased production of C. difficile toxins A and B in a defined medium by Yamakawa et al.,40,41 but not by Karlsson et al.42 If it is assumed that biotin-limited conditions may suppress toxin production, then a reduction in biotin-producing bifidobacteria could also have an impact upon toxin production. Nutritional effects of C. difficile toxin production have not been investigated during this experiment, but the gut model provides scope for such investigations to be carried out in a controlled, gut-reflective manner.
In contrast, few reports have associated the presence of Bacteroides spp. with colonization resistance.16,33 The marked effect of instillation of a combination of CTX/dCTX on Bacteroides spp., in comparison with CTX alone, may help to explain why previous test-tube experiments failed to explain the propensity of CTX towards C. difficile infection, and postulates a role for these organisms in colonization resistance. The considerable deleterious effect of CTX/dCTX upon Bacteroides spp. may, in turn, lead to liberation of endotoxin, which may exacerbate the inflammatory response of the gut mucosa provoked by C. difficile toxins. While the data presented here do not provide a definitive answer as to which components of gut flora are responsible for colonization resistance, they may indicate a prominent role for Bacteroides spp. and, to a lesser extent, bifidobacteria. These results allow further, more detailed investigations of these putative colonization resistance components.
C. difficile appeared to respond to antibiotic exposure by germinating and producing toxin in V2 and V3. Reports on antibiotic stimulation of toxin production in C. difficile are somewhat contradictory,4345 and in-house results demonstrate no difference in toxin production between C. difficile exposed to antibiotic and non-exposed controls. In contrast, the gut model clearly demonstrates C. difficile stasis in the absence of antibiotic pressure, followed by activation of the organism, leading to toxin production, after CTX or CTX/dCTX exposure. Previous in-house experiments showed that the UK epidemic strain produced a peak toxin titre of 56, regardless of exposure to antibiotics. The present study demonstrates peak titres of 5 during antibiotic instillation, suggesting that the increases in toxin titres were simply due to normal toxin production by the now activated C. difficile, as opposed to enhanced toxin release due to antibiotic stimulus. The delay in toxin production by C. difficile indicates that it may occur following an initial effect upon gut flora components, rather than a direct effect upon C. difficile itself. This may explain why CDI does not occur within hours of antibiotic administration, rather days or even weeks later.
Whereas the C. difficile population in V1 also appeared to germinate and proliferate, there was little evidence of the presence of toxin. It is not clear whether toxin production itself was repressed, or whether toxin was produced but was rapidly degraded at the low pH of this vessel. Low pH has previously been associated with low levels of toxin in some studies,15,46 but not in others.47 A recent report described acid-induced conformational changes in C. difficile toxin B. It is not known whether a shift to neutral pH causes refolding to the original toxin B conformation, but this may explain the apparent discrepancies in earlier results by way of methodological variation.47 Alternatively, it may be that C. difficile cytotoxin production is suppressed by acidic conditions. Further work at the level of gene expression is required alongside biological cytotoxin assays, to ascertain whether this is the case. There seemed little difference in the behaviour of C. difficile toxin production when exposed to the two antibiotic modes, although germination and cell growth was much more rapid following CTX/dCTX in V2 and V3. This may have been due to the residual effects of exposure to CTX alone, allowing a faster C. difficile response to combination instillation in these two vessels. It would be prudent to examine the effects of the two treatments when administered in reverse order, to ascertain whether this faster response is a true phenomenon, or due to residual antibiotic effects. In vivo studies of the effects of antibiotic treatment upon intestinal microorganisms suggest that recovery of bowel flora takes place within 12 weeks following cessation of treatment.48,49 While the present study used a 7 day recovery period, further investigations incorporating extended recovery periods may also be necessary. The exposure of the gut flora to antimicrobials may affect its recovery, by allowing the selection of antibiotic-resistant populations. This was not investigated in this study (owing to limited amounts of dCTX), and while overall populations seemed to recover well, the proportions of resistant species within those populations are unknown.
The possible contribution of dCTX to the propensity of CTX treatment to predispose towards CDI has not been examined previously. This is surprising considering the well-documented synergic effects of CTX/dCTX against anaerobes, and in particular, Bacteroides spp.2123 The present study demonstrates a clear depletion of Bacteroides spp. in V1 by CTX/dCTX over and above that achieved by CTX alone, although at a higher total drug exposure of CTX/dCTX, and is reflected in less marked population decreases in V2 and V3. This may explain why in vitro studies have so far failed to demonstrate predisposition to CDI by CTX. The results also indicate a similar, although less profound, effect upon bifidobacteria, which requires further investigation. Although CTX exposure alone resulted in C. difficile proliferation and toxin production, depletion of gut bacteria was less marked. Pharmacokinetic studies of CTX metabolism and elimination in elderly patients showed that elimination of the drug decreased with age of the patient.50 The intestinal flora of elderly patients may therefore be subjected to prolonged exposure to CTX, and by inference its metabolite dCTX, compared with that seen in younger patients.
The study of the interplay between antimicrobials, C. difficile and the intestinal microflora has so far relied upon the hamster model,5154 or upon single sample assays of faecal bacterial populations in volunteers.33,39,55 Such approaches are extremely difficult to control in terms of antibiotic exposure of bowel flora and C. difficile challenge. Following treatment with piperacillintazobactam 4.5 g at a ratio of 8:1, four of 17 faecal specimens had quantifiable levels of both piperacillin and tazobactam, the ratios of piperacillin:tazobactam being 1.8, 6.4, 7.6 and 11.1.19 This illustrates the wide variation in antibiotic concentrations achieved in vivo, and the difficulty in gaining reproducible results from investigations using human volunteers. In contrast, this gut model potentially provides a system that better reflects the conditions experienced in vivo, while allowing controlled exposure to both antibiotic and C. difficile. Whereas additional work using comparator antibiotics that do not predispose to CDI is necessary in order to validate this system further, as are further experiments to assess reproducibility of the findings, it represents a promising method for studying the pathogenesis of CDI.
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Acknowledgements |
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Footnotes |
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References |
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2 . George, W. L., Rolfe, R. D., Harding, G. K. et al. (1982). Clostridium difficile and cytotoxin in feces of patients with antimicrobial agent-associated pseudomembranous colitis. Infection 10, 2058.[ISI][Medline]
3 . Simor, A. E., Yake, S. L. & Tsimidis, K. (1993). Infection due to Clostridium difficile among elderly residents of a long-term-care facility. Clinical Infectious Diseases 17, 6728.[ISI][Medline]
4 . Climo, M. W., Israel, D. S., Wong, E. S. et al. (1998). Hospital-wide restriction of clindamycin: effect on the incidence of Clostridium difficile-associated diarrhea and cost. Annals of Internal Medicine 128, 98995.
5
.
Pear, S. M., Williamson, T. H., Bettin, K. M. et al. (1994). Decrease in nosocomial Clostridium difficile-associated diarrhea by restricting clindamycin use. Annals of Internal Medicine 120, 2727.
6 . de Lalla, F., Privitera, G., Ortisi, G. et al. (1989). Third generation cephalosporins as a risk factor for Clostridium difficile-associated disease: a four-year survey in a general hospital. Journal of Antimicrobial Chemotherapy 23, 62331.[Abstract]
7
.
Impallomeni, M., Galletly, N. P., Wort, S. J. et al. (1995). Increased risk of diarrhoea caused by Clostridium difficile in elderly patients receiving cefotaxime. British Medical Journal 311, 13456.
8 . Golledge, C. L., McKenzie, T. & Riley, T. V. (1989). Extended spectrum cephalosporins and Clostridium difficile. Journal of Antimicrobial Chemotherapy 23, 92931.[Abstract]
9 . Settle, C. D., Wilcox, M. H., Fawley, W. N. et al. (1998). Prospective study of the risk of Clostridium difficile diarrhoea in elderly patients following treatment with cefotaxime or piperacillin-tazobactam. Alimentary Pharmacological Therapy 12, 121723.[CrossRef]
10 . Bates, C. J., Wilcox, M. H., Spencer, R. C. et al. (1990). Ciprofloxacin and Clostridium difficile infection. Lancet 336, 1193.
11 . Hillman, R. J., Rao, G. G., Harris, J. R. et al. (1990). Ciprofloxacin as a cause of Clostridium difficile-associated diarrhoea in an HIV antibody-positive patient. Journal of Infection 21, 2057.[ISI][Medline]
12 . Anonymous. (1990). Ciprofloxacin and pseudomembranous colitis. Lancet 336, 150910.
13 . Cain, D. B. & OConnor, M. E. (1990). Pseudomembranous colitis associated with ciprofloxacin. Lancet 336, 946.[ISI][Medline]
14 . Golledge, C. L., Carson, C. F., ONeill, G. L. et al. (1992). Ciprofloxacin and Clostridium difficile-associated diarrhoea. Journal of Antimicrobial Chemotherapy 30, 1417.[Abstract]
15 . Borriello, S. P. & Barclay, F. E. (1986). An in-vitro model of colonisation resistance to Clostridium difficile infection. Journal of Medical Microbiology 21, 299309.[Abstract]
16 . Ambrose, N. S., Johnson, M., Burdon, D. W. et al. (1985). The influence of single dose intravenous antibiotics on faecal flora and emergence of Clostridium difficile. Journal of Antimicrobial Chemotherapy 3, 31926.
17 . Rolfe, R. D., Helebian, S. & Finegold, S. M. (1981). Bacterial interference between Clostridium difficile and normal fecal flora. Journal of Infectious Diseases 143, 4705.[ISI][Medline]
18 . Freeman, J. & Wilcox, M. H. (2000). Does antibiotic exposure enhance sporulation of an epidemic Clostridium difficile strain? In Program and Abstracts of the Fortieth Interscience Conference on Antimicrobial Agents and Chemotherapy, Toronto, Ontario, Canada, 2000. Abstract 950, p. 412. American Society for Microbiology, Washington, DC, USA.
19
.
Wilcox, M. H., Brown, A. & Freeman, J. (2001). Faecal concentrations of piperacilllin and tazobactam in elderly patients. Journal of Antimicrobial Chemotherapy 48, 1556.
20 . Novick, W. J. (1982). Levels of cefotaxime in body fluids and tissues: a review. Reviews of Infectious Diseases 4, S34653.[ISI][Medline]
21 . Jones, R. N. (1995). Cefotaxime and desacetylcefotaxime antimicrobial interactions. The clinical relevance of enhanced activity: a review. Diagnostic Microbiology and Infectious Diseases 22, 1933.[CrossRef][ISI][Medline]
22 . Canawati, H. N. (1992). A reassessment of the activity of the third-generation cephalosporins against anaerobes and Staphylococcus aureus. American Journal of Surgery 164 24S7S.[Medline]
23 . Aldridge, K. E. (1989). Comparison of the in vitro action and interaction of cefotaxime and desacetylcefotaxime against clinical isolates of Bacteroides spp. Diagnostic Microbiology and Infectious Diseases 12, 4550.[ISI][Medline]
24 . Guggenbichler, J. P., Kofler, J. & Allerberger, F. (1985). The influence of third-generation cephalosporins on the aerobic intestinal flora. Infection 13, Suppl. 1, S1279.
25 . Knothe, H., Dette, A. H. & Shah, P. M. (1985). Impact of injectable cephalosporins on the gastrointestinal microflora: observations in healthy volunteers and hospitalised patients. Infection 13, Suppl. 1, S12933.[ISI][Medline]
26 . Macfarlane, G. T., Macfarlane, S. & Gibson, G. R. (1998). Validation of a three-stage compound continuous culture system for investigating the effect of retention time on the ecology and metabolism of bacteria in the human colon. Microbial Ecology 35, 1807.[CrossRef][ISI][Medline]
27 . McBain, A. J. & MacFarlane, G. T. (1998). Ecological and physiological studies on large intestinal bacteria in relation to production of hydrolytic and reductive enzymes involved in formation of genotoxic metabolites. Journal of Medical Microbiology 47, 40716.[Abstract]
28 . Olano-Martin, E., Mountzouris, K. C., Gibson, G. R. et al. (2000). In vitro fermentability of dextran, oligodextran and maltodextrin by human gut bacteria. British Journal of Nutrition 83, 24755.[ISI][Medline]
29
.
Stubbs, S. L., Brazier, J. S., ONeill, G. L. et al. (1999). PCR targeted to the 16S-23S rRNA gene intergenic spacer region of Clostridium difficile and construction of a library consisting of 116 different PCR ribotypes. Journal of Clinical Microbiology. 37, 4613.
30 . Ebright, J. R., Fekety, R., Silva, J. et al. (1981). Evaluation of eight cephalosporins in hamster colitis model. Antimicrobial Agents and Chemotherapy 19, 9806.[ISI][Medline]
31 . Larson, H. E. & Welch, A. (1993). In-vitro and in-vivo characterisation of resistance to colonisation with Clostridium difficile. Journal of Medical Microbiology 38, 1038.[Abstract]
32 . Onderdonk, A. B., Brodasky, T. F. & Bannister, B. (1981). Comparative effects of clindamycin and clindamycin metabolites in the hamster model of antibiotic-associated colitis. Journal of Antimicrobial Chemotherapy 8, 38393.[ISI][Medline]
33
.
Hopkins, M. J. & Macfarlane, G. T. (2002). Changes in predominant bacterial populations in human faeces with age and with Clostridium difficile infection. Journal of Medical Microbiology 51, 44854.
34
.
Hopkins, M. J., Sharp, R. & Macfarlane, G. T. (2001). Age and disease related changes in intestinal bacterial populations assessed by cell culture, 16S rRNA abundance, and community cellular fatty acid profiles. Gut 48, 198205.
35 . Ouwehand, A. C., Isolauri, E., Kirjavainen, P. V. et al. (1999). Adhesion of four Bifidobacterium strains to human intestinal mucus from subjects in different age groups. FEMS Microbiology Letters 172, 614.[CrossRef][ISI][Medline]
36 . Wang, X. & Gibson, G. R. (1993). Effects of the in vitro fermentation of oligofructose and inulin by bacteria growing in the human large intestine. Journal of Applied Bacteriology 75, 37380.[ISI][Medline]
37 . Gibson, G. R. & Wang, X. (1994). Regulatory effects of bifidobacteria on the growth of other colonic bacteria. Journal of Applied Bacteriology 77, 41220.[ISI][Medline]
38 . Benno, Y. & Mitsuoka, T. (1992). Impact of Bifidobacterium longum on human fecal microflora. Microbiology and Immunology 36, 68394.[ISI][Medline]
39 . Kim, S. H., Yang, S. J. & Koo, H. C. et al. (2001). Inhibitory activity of Bifidobacterium longum HY8001 against Vero cytotoxin of Escherichia coli O157:H7. Journal of Food Protection 64, 166773.[ISI][Medline]
40 . Yamakawa, K., Karasawa, T., Ikoma, S. et al. (1996). Enhancement of Clostridium difficile toxin production in biotin-limited conditions. Journal of Medical Microbiology 44, 11114.[Abstract]
41 . Yamakawa, K., Karasawa, T., Ohta, T. et al. (1998). Inhibition of enhanced toxin production by Clostridium difficile in biotin-limited conditions. Journal of Medical Microbiology 47, 76771.[Abstract]
42 . Karlsson, S., Burman, L. G. & Akerlund, T. (1999). Suppression of toxin production in Clostridium difficile VPI 10463 by amino acids. Microbiology 145, 168393.[Abstract]
43 . Barc, M. C., Depitre, C., Corthier, G. et al. (1992). Effects of antibiotics and other drugs on toxin production in Clostridium difficile in vitro and in vivo. Antimicrobial Agents and Chemotherapy 36, 13326.[Abstract]
44 . Honda, T., Hernadez, I., Katoh, T. et al. (1983). Stimulation of enterotoxin production of Clostridium difficile by antibiotics. Lancet 1, 655.
45 . Nakamura, S., Mikawa, M., Tanabe, N. et al. (1982). Effect of clindamycin on cytotoxin production by Clostridium difficile. Microbiology and Immunology 26, 98592.[Medline]
46 . Onderdonk, A. B., Lowe, B. R. & Bartlett, J. G. (1979). Effects of environmental stress on Clostridium difficile toxin levels during continuous cultivation. Applied and Environmental Microbiology 38, 63741.[ISI][Medline]
47
.
QaDan, M., Spyres, L. M. & Ballard, J. D. (2000). pH-induced conformational changes in Clostridium difficile toxin B. Infection and Immunity 68, 24704.
48 . Adamsson, I., Edlund, C., Sjostedt, S. et al. (1997). Comparative effects of cefadroxil and phenoxymethylpenicillin on the normal oropharyngeal and intestinal microflora. Infection 25, 1548.[ISI][Medline]
49 . Lode, H., Muller, C., Borner, K. et al. (1992). Multiple-dose pharmacokinetics of cefprozil and its impact on intestinal flora of volunteers. Antimicrobial Agents and Chemotherapy 36, 1449.[Abstract]
50 . Ludwig, E., Szekely, E., Csiba, A. et al. (1988). Pharmacokinetics of cefotaxime and desacetylcefotaxime in elderly patients. Drugs 35, Suppl. 2, 516.
51 . Borriello, S. P. & Barclay, F. E. (1985) Protection of hamsters against Clostridium difficile ileocaecitis by prior colonisation with non-pathogenic strains. Journal of Medical Microbiology 19, 33950.[Abstract]
52 . Borriello, S. P., Ketley, J. M., Mitchell, T. J. et al. (1987). Clostridium difficilea spectrum of virulence and analysis of putative virulence determinants in the hamster model of antibiotic-associated colitis. Journal of Medical Microbiology 24, 5364.[Abstract]
53 . Borriello, S. P., Welch, A. R., Barclay, F. E. et al. (1988). Mucosal association by Clostridium difficile in the hamster gastrointestinal tract. Journal of Medical Microbiology 25, 1916.[Abstract]
54 . Larson, H. E. & Borriello, S. P. (1990). Quantitative study of antibiotic-induced susceptibility to Clostridium difficile enterocecitis in hamsters. Antimicrobial Agents and Chemotherapy 34, 134853.[ISI][Medline]
55 . Nord, C. E. & Heimdahl, A. (1986). Impact of orally administered antimicrobial agents on human oropharyngeal and colonic microflora. Journal of Antimicrobial Chemotherapy 18, Suppl. C, 15964.[Abstract]