SUPPLEMENT ARTICLE Antibiotic Regimens and Intestinal Colonization with Antibiotic-Resistant Gram-Negative Bacilli Curtis J. Donskey Infectious Diseases Section, Louis Stokes Cleveland Veterans Affairs Medical Center, Cleveland, Ohio The intestinal tract provides an important reservoir for antibiotic-resistant gram-negative bacilli, including Enterobacteriaceae species, Pseudomonas aeruginosa, and Acinetobacter species [1–9]. Although most patients who are colonized with these organisms remain asymptomatic, infections may occur because of translocation across the intestinal lining or as a consequence of fecal contamination of wounds or devices [2, 9]. Several studies have shown that intestinal colonization by gram-negative bacilli often precedes the onset of infection [1–4]. Fecal shedding onto patients’ skin and environmental surfaces contributes to nosocomial transmission of antibioticresistant gram-negative pathogens . Finally, the intestinal tract provides an important site for transfer of genes conferring antibiotic resistance . Selective pressure exerted by antibiotics plays a crucial role in the emergence and dissemination of antibiotic-resistant microorganisms. This review is an examination of the effects of antibiotic treatment on Reprints or correspondence: Dr. Curtis J. Donskey, Infectious Diseases Section, Louis Stokes Cleveland Veterans Affairs Medical Center, 10701 East Blvd., Cleveland, OH 44106 ([email protected]). Clinical Infectious Diseases 2006; 43:S62–9 This article is in the public domain, and no copyright is claimed. 1058-4838/2006/4305S2-0005 S62 • CID 2006:43 (Suppl 2) • Donskey colonization of the intestinal tract with antibioticresistant gram-negative bacilli. Findings from studies involving animal models and healthy human volunteers are used to illustrate general concepts regarding the effects of antibiotics on colonization with pathogens, and the applicability of these concepts to clinical settings and implications for control of antibiotic-resistant gram-negative bacilli are discussed. COLONIZATION RESISTANCE The indigenous bacteria of the colon provide an important host-defense mechanism by inhibiting colonization by potentially pathogenic microorganisms. This defense mechanism, termed “colonization resistance,” can be applied to the prevention of overgrowth by indigenous potential pathogens and the inhibition of colonization by exogenously introduced organisms . Escherichia coli, a member of the indigenous colonic microflora, is normally maintained at relatively low population densities by the predominant anaerobic microflora . Healthy humans ingesting small numbers of P. aeruginosa (102 cfu) do not develop detectable levels of organisms in stool; however, larger inocula (⭓106 cfu) have been found to result in shedding in stool for 1–6 days . Foods such as salads may contain relatively large numbers of P. aeruginosa or Entero- Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 The intestinal tract provides an important reservoir for antibiotic-resistant gram-negative bacilli, including Enterobacteriaceae species, Pseudomonas aeruginosa, and Acinetobacter baumannii. Selective pressure exerted by antibiotics plays a crucial role in the emergence and dissemination of these pathogens. Many classes of antibiotics may promote intestinal colonization by health care–associated gram-negative bacilli, because the organisms are often multidrug resistant. Antibiotics may inhibit colonization by gram-negative pathogens that remain susceptible, but the benefits of this effect are often limited because of the emergence of resistance. Antibiotic formulary alterations and standard infection control measures have been effective in controlling outbreaks of colonization and infection with antibiotic-resistant gram-negative pathogens. Additional research is needed to clarify the role of strategies such as selective decontamination of the digestive tract and decontamination of environmental surfaces and of patients’ skin and wounds. bacteriaceae species (103–104 cfu/serving) , a finding that could, in part, explain why P. aeruginosa has been detected in stool of patients with cancer who have not received prior antibiotic treatment [3, 14]. Experimental ingestion of exogenous E. coli by healthy humans does not typically result in persistent colonization [12, 15]; however, travelers to Mexico frequently acquire colonization by antibiotic-resistant strains of E. coli in the absence of prophylactic or therapeutic antibiotic treatment . Acquisition in this setting could be due to repeated ingestion of large numbers of organisms and/or special properties of the ingested organisms (e.g., the ability to adhere to the mucosa) . ANTIBIOTICS AND COLONIZATION WITH GRAM-NEGATIVE BACILLI Antibiotics and Gram-Negative Bacilli • CID 2006:43 (Suppl 2) • S63 Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 Antibiotic selective pressure. Bacteria possess a remarkable ability to develop and acquire resistance to antibiotics. Among gram-negative bacilli, common mechanisms of resistance include modification of drug targets; production of inactivating enzymes, such as b-lactamases and aminoglycoside-modifying enzymes; efflux pumps; and alterations in outer membrane proteins that are associated with decreased permeability to antibiotics . In general, antibiotic exposure is not thought to directly induce these resistance mechanisms. Rather, antibiotic therapy promotes proliferation of antibiotic-resistant gramnegative bacilli by exerting selective pressure (i.e., inhibition of competing microflora but not of resistant organisms). In individual patients, selective pressure may facilitate the emergence of new resistant mutants or of preexisting subpopulations of resistant organisms. For example, ceftazidime therapy may eliminate susceptible gram-negative bacilli while allowing expansion of the population of a new mutant of Klebsiella pneumoniae that harbors an extended-spectrum b-lactamase (ESBL) or of a preexisting subpopulation of Enterobacter species that constitutively hyperproduces chromosomal cephalosporinases [19–21]. Numerous clinical studies have documented the emergence of resistant gram-negative bacilli in association with the use of agents for treatment of gram-negative pathogens [4–7, 19–28]. Although this review focuses on the intestinal tract as a site for emergence of antibiotic-resistant gram-negative bacilli, resistant organisms also emerge frequently from other sites. Once antibiotic-resistant gram-negative pathogens have emerged, antibiotics play a crucial role in their subsequent spread from patient to patient . Antibiotic therapy may markedly reduce the number of exogenous bacteria that must be ingested to establish intestinal colonization [9, 11, 13]. Antibiotic-associated overgrowth of nosocomial pathogens and antibiotic-associated diarrhea contribute to increased shedding of organisms onto patients’ skin or environmental surfaces [29, 30]. Organisms on skin or surfaces may then be acquired on health care workers’ hands . Because health care–associated gram-negative bacilli are often resistant to multiple classes of antibiotics, many different antibiotics may potentially facilitate colonization and dissemination of these pathogens. For example, third-generation cephalosporins, trimethoprim-sulfamethoxazole, ciprofloxacin, and aminoglycosides have all been associated with ESBL-producing gram-negative bacilli [7, 31]. In an outbreak of colonization and infection with ESBL-producing gram-negative bacilli in nursing homes, most patients had not received prior ceftazidime . Rather, receipt of ciprofloxacin or trimethoprim-sulfamethoxazole was an independent risk factor for colonization; determinants of resistance to ceftazidime and trimethoprim-sulfamethoxazole were linked on a plasmid, whereas ciprofloxacin resistance was not directly linked to ceftazidime resistance . Similarly, piperacillin-tazobactam, imipenem, aminoglycosides, vancomycin, and broad-spectrum cephalosporins have all been associated with piperacillin- and tazobactam-resistant P. aeruginosa . Although antibiotics may promote proliferation of antibiotic-resistant gram-negative pathogens, these agents may also provide a protective effect if they have inhibitory activity [9, 33–36]. For example, Kaye et al.  found a trend toward reduced isolation of E. coli resistant to ampicillin and sulbactam in patients exposed to piperacillin-tazobactam, an agent with activity against many isolates that are resistant to ampicillin and sulbactam. In another study, fluoroquinolone exposure was found to be protective against isolation of broad-spectrum cephalosporin-resistant Enterobacter species. . Paterson et al.  used oral norfloxacin to inhibit intestinal colonization by fluoroquinolone-susceptible ESBL-producing E. coli during an outbreak among patients undergoing liver transplantation. However, increasing rates of fluoroquinolone resistance among gram-negative bacilli, including ESBL producers, is likely to limit the protective effect of these agents [37, 38]. As was noted above, prior receipt of ciprofloxacin has been shown to be an independent risk factor for colonization with ESBL-producing gram-negative bacilli resistant to fluoroquinolones . Animal models. Mouse models provide a useful means of directly comparing the effects of antibiotics on intestinal colonization with nosocomial pathogens. During treatment, antibiotics that are excreted into the intestinal tract may potentially inhibit gram-negative pathogens and competing indigenous microflora. After completion of treatment, the indigenous microflora recover over several days. Susceptibility to colonization by resistant gram-negative bacilli may persist during the recovery period. Figure 1 summarizes the findings of my group’s mouse model studies examining establishment of colonization by ESBL-producing K. pneumoniae [39–41] (au- thor’s unpublished data). ESBL-producing K. pneumoniae (104 cfu) was administered orally once during and once 2 days after completion of subcutaneous antibiotic treatment. Antibiotics that disrupted the anaerobic microflora and possessed minimal activity against the K. pneumoniae isolate (e.g., clindamycin and linezolid) promoted colonization. An antibiotic that disrupted the anaerobic microflora and possessed significant activity against the K. pneumoniae isolate (piperacillin-tazobactam [MIC, 4 mg/mL]) inhibited the establishment of colonization during treatment but promoted overgrowth when exposure occurred during the period of recovery of the indigenous microflora. Antibiotics that did not disrupt the anaerobic microflora (e.g., cefepime, aztreonam, levofloxacin, and daptomycin) did not promote colonization. These findings are very similar to findings from mouse studies that examined the effects of antibiotics on colonization by vancomycin-resistant enterococci (VRE) and Clostridium difficile [9, 42]. Of note, however, Hentges et al.  found that oral streptomycin promoted Pseudomonas aeruginosa intestinal colonization and translocation across the intestinal lining to a greater degree than did oral clindamycin; oral streptomycin inhibits facultative anaerobes and obligate anaerobes, whereas clindamycin selectively inhibits anaerobes. Although animal models have limitations, these studies may raise important issues that deserve further study in patients. First, animal studies may implicate antibiotics, such as clindamycin and linezolid, that have not been associated with anS64 • CID 2006:43 (Suppl 2) • Donskey Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 Figure 1. Effect of antibiotic treatment on establishment of intestinal colonization with extended-spectrum b-lactamase–producing Klebsiella pneumoniae in mice. Mice received subcutaneous antibiotic treatment from day ⫺2 to day 3 (solid bar), and oral extended-spectrum b-lactamase–producing K. pneumoniae (10,000 cfu) was administered once during treatment and once 2 days after completion of treatment. Densities of extended-spectrum b-lactamase–producing K. pneumoniae in stool are shown. If extended-spectrum b-lactamase–producing K. pneumoniae were not detected, the lower limit of detection was assigned (2 log10 cfu/g). Pip, piperacillin. tibiotic-resistant gram-negative bacilli in clinical studies. It is plausible that these antibiotics may promote overgrowth of gram-negative pathogens in patients, because both promote overgrowth of Enterobacteriaceae species in healthy humans and because clindamycin promotes the emergence of new gram-negative bacilli . Second, the promotion of piperacillin- and tazobactam-susceptible K. pneumoniae by piperacillin-tazobactam in mice exposed after the treatment period (figure 1) illustrates that adverse effects of antibiotics often extend beyond the period of treatment. Although recovery of the microflora of mice or healthy humans occurs within 1–2 weeks after antibiotic therapy is stopped , longer delays in recovery of protective indigenous microflora may occur in patients who have received multiple or prolonged courses of antibiotics. Finally, animal model studies suggest that antibiotics that do not disrupt the anaerobic microflora may be less likely to promote colonization by resistant gram-negative bacilli [39– 41]. More data are needed to evaluate whether selective use of such agents may offer any advantage to patients. Healthy volunteers. Many studies have been performed to evaluate the effect of different antibiotics on the indigenous intestinal microflora of humans . Most studies have been performed in healthy volunteers, and clinical studies usually have involved monotherapy regimens in patients with mild to moderate severity of illness. As has been noted previously, these studies demonstrate that antibiotics that inhibit anaerobes without inhibiting Enterobacteriaceae species (e.g., clindamycin, linezolid, and oral vancomycin) may promote overgrowth of indigenous Enterobacteriaceae species and the emergence of new antibiotic-resistant gram-negative bacilli . However, antibiotics that cause relatively little disruption of the anaerobic microflora (e.g., trimethoprim-sulfamethoxazole, cefadroxil, and ciprofloxacin) have also been shown to promote the emergence of antibiotic-resistant gram-negative bacilli [11, 44]. Although studies of healthy volunteers provide a useful reference regarding the potential impact of antibiotics on colonization with gram-negative bacilli, several factors may limit their applicability to clinical settings. First, patients are at increased risk of exposure to exogenous antibiotic-resistant gramnegative bacilli. The oropharynx of a hospitalized patient frequently becomes colonized with gram-negative bacilli that may be ingested. Nasogastric tubes may facilitate colonization of the oropharynx by P. aeruginosa that form biofilms on plastic surfaces . Second, patients often receive proton pump inhibitors or histamine2 blockers that inhibit production of stomach acid . These agents promote overgrowth of gram-negative bacilli in the stomach and facilitate passage of organisms into the small intestine . Third, the colonic microflora of ill hospitalized patients may be altered in the absence of antibiotic therapy . Finally, patients often receive multiple antibiotics Figure 2. Densities of indigenous and acquired facultative gram-negative bacilli in stool samples from 5 healthy volunteers receiving treatment with oral ciprofloxacin (20 mg/day) for 14 days followed by oral ciprofloxacin in combination with clindamycin (300 mg/day). Ciprofloxacin monotherapy eliminated indigenous Escherichia coli (dotted lines). No subjects acquired exogenous ciprofloxacin-resistant gram-negative bacilli during ciprofloxacin monotherapy, whereas 3 subjects did during the combination treatment period (solid circles). The acquired exogenous gram-negative bacilli included E. coli (2 strains) and Citrobacter freundii. Data are from Joris et al. . Effect of antibiotics with activity against intestinal anaerobes. Antibiotics that inhibit intestinal anaerobes promote overgrowth of VRE in stools of colonized patients . We tested the hypothesis that such antibiotic regimens may also promote overgrowth of coexisting gram-negative bacilli resistant to ceftazidime, ciprofloxacin, or piperacillin-tazobactam in stool of patients colonized with VRE . As shown in figure 3A and 3B, therapy with antianaerobic antibiotic regimens was associated with an increased likelihood that an antibioticresistant gram-negative bacillus would be isolated, and, when present, the density of these organisms was higher during therapy than in the absence of antianaerobic therapy for at least 2 weeks. These findings suggest that efforts to limit the use of antianaerobic antibiotics could minimize the density of VRE and coexisting antibiotic-resistant gram-negative bacilli. However, it should be noted that many antibiotics with antianaerobic activity also have activity against gram-negative bacilli. For example, figure 3C shows the emergence of an isolate of E. coli resistant to piperacillin-tazobactam in stool of a patient during treatment with piperacillin-tazobactam, followed by elimination of the organism during meropenem therapy . CONTROL STRATEGIES FOR ANTIBIOTICRESISTANT GRAM-NEGATIVE BACILLI Infection control measures. Standard infection control measures play a crucial role in limiting the spread of antibioticresistant gram-negative bacilli. Because the hands of health care workers often become contaminated with gram-negative bacilli, efforts to improve adherence to hand hygiene are essential . Alcohol-based hand-hygiene products are effective at eliminating gram-negative bacilli, and the use of these agents in combination with ongoing education has been associated with reductions in nosocomial infections . For organisms resistant to multiple antibiotics or during outbreaks, contact precautions may be indicated . Surveillance for stool carriage of antibiotic-resistant gram-negative bacilli may be helpful in some situations, because colonized patients often outnumber those with clinical isolates [6, 9]. Lucet et al.  used a multifaceted infection control intervention with no concurrent antibiotic formulary changes to control high rates of colonization and infection with endemic ESBL-producing organisms. Because multiple nosocomial pathogens resistant to antibiotics often coexist in the same patient populations , efforts to improve infection control practices may offer the additional benefit of limiting the spread of coexisting pathogens [6, 9, 51]. Reducing the burden of antibiotic-resistant gram-negative pathogens present on patients’ skin and on environmental surfaces might potentially reduce transmission by decreasing the number of organisms acquired on health care workers’ hands and by decreasing direct transmission from surfaces to patients . A recent study found that patients harboring antibioticAntibiotics and Gram-Negative Bacilli • CID 2006:43 (Suppl 2) • S65 Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 concurrently or in sequence, which makes it difficult to determine the effects of individual agents. One illustration of the discrepancy that may occur between healthy volunteers and patients is provided by studies of fluoroquinolone antibiotics. In healthy volunteers, most fluoroquinolones inhibit Enterobacteriaceae species but cause minimal disruption of intestinal anaerobes, and acquisition of fluoroquinolone-resistant gram-negative bacilli is uncommon [44, 47]. In contrast, numerous clinical studies have demonstrated that fluoroquinolone use may be associated with fluoroquinolone-resistant gram-negative pathogens [26, 28, 37, 38]. In addition to being at increased risk for exposure to fluoroquinolone-resistant gram-negative bacilli, hospitalized patients may have preexisting colonization with such organisms, which may expand in population during fluoroquinolone therapy. The fact that hospitalized patients often receive fluoroquinolones in combination with other antibiotics may contribute significantly to the potential for acquisition and overgrowth of fluoroquinolone-resistant gram-negative organisms. Joris et al.  illustrated this by giving low-dose ciprofloxacin monotherapy to healthy outpatient volunteers, followed by ciprofloxacin in combination with oral clindamycin. As shown in figure 2, no ciprofloxacin-resistant gram-negative bacilli were detected during ciprofloxacin monotherapy, but 3 of 5 subjects acquired new ciprofloxacin-resistant gram-negative bacilli when clindamycin was added . S66 • CID 2006:43 (Suppl 2) • Donskey Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 Figure 3. Effect of anti-anaerobic antibiotic regimens on the detection and density of antibiotic-resistant gram-negative bacilli in stool of patients colonized with vancomycin-resistant enterococci (VRE). A, Detection of gram-negative bacilli resistant to ceftazidime, ciprofloxacin, or piperacillintazobactam in stool of patients during therapy with anti-anaerobic antibiotic regimens and in the absence of such therapy for 11 month. B, Density of resistant gram-negative bacilli in stool during anti-anaerobic antibiotic therapy and in the absence of such therapy for 12 weeks (among patients with detectable antibiotic-resistant gram-negative bacilli). C, Effect of antibiotic therapy on the density of VRE (triangles) and piperacillin-tazobactam–resistant Escherichia coli (squares) in the stool of a patient. Antibiotic therapy included piperacillin-tazobactam, vancomycin, and metronidazole (regimen A); piperacillin-tazobactam and vancomycin (regimen B); and meropenem and linezolid (regimen C). The star indicates development of catheter-related bacteremia with an E. coli isolate with a susceptibility pattern identical to that of the stool isolate. Treatment with meropenem and linezolid (regimen C) resulted in suppression of the 2 pathogens to undetectable levels in stool (reprinted with permission from ). resistant gram-negative pathogens were less likely to have environmental contamination than were those harboring resistant gram-positive pathogens (4.9% vs. 24.7%) . However, extensive environmental contamination with Acinetobacter baumannii has been described, and this organism survives for prolonged periods on surfaces . In addition, P. aeruginosa may persistently contaminate moist areas, such as sinks . Several quasi-experimental studies have suggested that environmental decontamination may be a useful adjunctive measure for control of these pathogens [53–58]. Borer et al.  found that using chlorhexidine to disinfect the skin of patients in intensive care units was an effective means of reducing skin contamination with A. baumannii. Similarly, Urban et al.  used polymyxin B to decontaminate wounds as an adjunctive control measure for A. baumannii. Selective decontamination of the digestive tract. The goal of selective decontamination is to inhibit pathogens in the gastrointestinal tract without disturbing the anaerobic microflora . Typically, nonabsorbed antibiotics are applied to the oropharynx and ingested orally with or without concurrent administration of intravenous antibiotics. In a recent study in the Netherlands, a selective decontamination regimen including parenteral cefotaxime and oropharyngeal and enteral tobramycin, colistin, and amphotericin was associated with a reduction in ventilator-associated pneumonia and mortality among patients in an intensive care unit . Acceptance of selective decontamination in the United States has been limited, in part by concerns that the antibiotic regimens may promote overgrowth and infections with pathogens that are resistant to the agents being administered (e.g., VRE or methicillin-resistant Staphylococcus aureus) . Jackson et al.  have shown that the selective decontamination regimen used in the Dutch study causes overgrowth and translocation of indigenous enterococci in rats. Therefore, caution is indicated if these regimens are to be used in settings in which VRE and methicillin-resistant S. aureus are endemic. As has been noted elsewhere, oral norfloxacin has been used to selectively decontaminate intestinal colonization by fluoroquinolone-susceptible ESBL-producing organisms . Finally, oropharyngeal decontamination alone has been effective in preventing ventilator-associated pneumonia, suggesting that the combination of intravenous and orally ingested components of selective decontamination may not be necessary for prevention of this condition . Measures to control antibiotic use. One strategy for the control of antibiotic-resistant gram-negative bacilli is to limit antibiotic use, in an effort to reduce antibiotic selective pressure. In a teaching hospital in Cleveland, we found that 30% of days of antibiotic therapy among patients who were not in intensive care were unnecessary, on the basis of standard guidelines or standard principles of infectious diseases . These data demonstrate CONCLUSION Antibiotic selective pressure has contributed to the emergence and spread of antibiotic-resistant gram-negative pathogens. The intestinal tract provides an important reservoir for dissemination of these pathogens. Adherence to standard infection control measures and good antibiotic stewardship are essential control strategies. Additional research is needed to clarify the potential utility of strategies such as selective decontamination of the digestive tract and decontamination of environmental surfaces and of patients’ skin and wounds. Future directions for research should include efforts to develop novel technologies for control of antibiotic-resistant gram-negative pathogens. For example, we have demonstrated that oral administration of b-lactamase enzymes in conjunction with parenteral b-lactam antibiotics can degrade the portion of antibiotic that is excreted into the intestinal tract of mice, thereby preserving resistance against colonization by ESBL-producing K. pneumoniae . Others have developed light-activated antimicrobial coatings for continuous disinfection of surfaces . Acknowledgments I thank Robert Bonomo and Marion Helfand for critical manuscript review. Financial support. This work was supported by an Advanced Research Career Development Award from the Department of Veterans Affairs to C.J.D. Potential conflicts of interest. C.J.D. has received research grant support from Elan, Merck, Cubist, and Ortho-McNeil and serves on the speakers’ bureau of Elan and Ortho-McNeil. He is a consultant for Optimer Pharmaceuticals. References 1. Schimpff SC, Young VM, Greene WH, Vermeulen GD, Moody MR, Wiernik PH. Origin of infection in acute nonlymphocytic leukemia: significance of hospital acquisition of potential pathogens. Ann Intern Med 1972; 77:707–14. 2. Tancrede CH, Andremont AO. Bacterial translocation and gram-negative bacteremia in patients with hematological malignancies. J Infect Dis 1985; 152:99–103. 3. Wingard JR, Dick J, Charache P, Saral R. Antibiotic-resistant bacteria in surveillance stool cultures of patients with prolonged neutropenia. Antimicrob Agents Chemother 1986; 30:435–9. 4. Pena C, Pujol M, Ardanuy C, et al. Epidemiology and successful control of a large outbreak due to Klebsiella pneumoniae producing extendedspectrum beta-lactamases. Antimicrob Agents Chemother 1998; 42: 53–8. 5. Olson B, Weinstein RA, Nathan C, Chamberlin W, Kabins SA. Epidemiology of endemic Pseudomonas aeruginosa: why infection control efforts have failed. J Infect Dis 1984; 150:808–16. 6. Lucet J, Decre D, Fichelle A, et al. Control of a prolonged outbreak of extended-spectrum b-lactamase–producing Enterobacteriaceae in a university hospital. Clin Infect Dis 1999; 29:1411–8. 7. Weiner J, Quinn JP, Bradford PA, et al. Multiple antibiotic-resistant Klebsiella and Escherichia coli in nursing homes. JAMA 1999; 281: 517–23. 8. Corbella X, Pujol M, Ayats J, et al. Relevance of digestive tract colonization in the epidemiology of nosocomial infections due to multiresistant Acinetobacter baumannii. Clin Infect Dis 1996; 23:329–34. 9. Donskey CJ. The role of the intestinal tract as a reservoir and source for transmission of nosocomial pathogens. Clin Infect Dis 2004; 39: 219–26. 10. Salyers AA, Gupta A, Wang Y. Human intestinal bacteria as reservoirs for antibiotic resistance genes. Trends Microbiol 2004; 12:412–6. 11. Vollaard EJ, Clasener HA. Colonization resistance. Antimicrob Agents Chemother 1994; 38:409–14. 12. Freter R, Brickner H, Fekete J, Vickerman MM, Carey KE. Survival and implantation of Escherichia coli in the intestinal tract. Infect Immun 1983; 39:686–703. 13. Buck AC, Cooke EM. The fate of ingested Pseudomonas aeruginosa in normal persons. J Med Microbiol 1969; 2:521–5. 14. Remington JS, Schimpff SC. Please don’t eat the salads. N Engl J Med 1981; 304:433–5. 15. Cooke EM, Hettiaratchy IG, Buck AC. Fate of ingested Escherichia coli in normal persons. J Med Microbiol 1972; 5:361–9. 16. Murray BE, Mathewson JJ, Dupont HL, Ericsson CD, Reves RR. Emergence of resistant fecal Escherichia coli in travelers not taking prophylactic antimicrobial agents. Antimicrob Agents Chemother 1990; 34: 515–8. 17. Kaper JB, Nataro JP, Mobley HL. Pathogenic Escherichia coli. Nat Rev Microbiol 2004; 2:123–40. 18. Rice LB, Bonomo RA. Genetic and biochemical mechanisms of bacterial resistance to antimicrobial agents. In: Lorian V, ed. Antibiotics in laboratory medicine. 5th ed. Philadelphia: Lippincott Williams & Wilkins; 2005:441–508. Antibiotics and Gram-Negative Bacilli • CID 2006:43 (Suppl 2) • S67 Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 that significant reductions in antibiotic use in hospitals may be feasible. In addition, efforts to reduce the duration of antibiotic therapy have yielded promising results. For example, Chastre et al.  found that therapy for 8 days was as effective as therapy for 15 days for treatment of ventilator-associated pneumonia, and, among patients who developed recurrent infections, multidrug-resistant pathogens emerged less frequently in those who had received antibiotics for 8 days. Another strategy to control antibiotic-resistant gram-negative bacilli is the implementation of formulary changes that involve restricting the use of antibiotics that have frequently been associated with particular pathogens. Third-generation cephalosporins have often been targeted for restriction, because they have been associated with ESBL-producing gram-negative bacilli as well as with multidrug-resistant P. aeruginosa and Acinetobacter species [32, 58, 65]. Substitution of piperacillintazobactam, cefepime, imipenem, or ticarcillin-clavulanate for third-generation cephalosporins has been associated with reductions in ESBL-producing organisms . The rationale for such formulary alterations is that selective pressure due to third-generation cephalosporins is removed, and the other agents may suppress these pathogens in the intestinal tract or at other sites [9, 65]. Restriction of ciprofloxacin has also been associated with a decrease in ciprofloxacin resistance in P. aeruginosa isolates . Unfortunately, substitution of one agent for another may lead to the emergence of resistance to the new agent . Finally, it is noteworthy that the data supporting formulary alterations come primarily from quasi-experimental studies that should be interpreted cautiously because they are subject to a number of biases . S68 • CID 2006:43 (Suppl 2) • Donskey 38. Neuhauser MM, Weinstein RA, Rydman R, Danziger LH, Karam G, Quinn JP. Antibiotic resistance among gram-negative bacillus in US intensive care units: implications for fluoroquinolone use. JAMA 2003; 289:885–8. 39. Hoyen CK, Pultz NJ, Paterson DL, Aron DC, Donskey CJ. Effect of parenteral antibiotic administration on establishment of intestinal colonization in mice by Klebsiella pneumoniae strains producing extendedspectrum b-lactamases. Antimicrob Agents Chemother 2003; 47:3610–2. 40. Stiefel U, Pultz NJ, Helfand MS, Donskey CJ. Increased susceptibility to establishment of vancomycin-resistant Enterococcus intestinal colonization persists after completion of antianaerobic antibiotic treatment in mice. Infect Control Hosp Epidemiol 2004; 25:373–9. 41. Pultz NJ, Stiefel U, Donskey CJ. Effects of daptomycin, linezolid, and vancomycin on establishment of intestinal colonization with vancomycin-resistant enterococci and extended-spectrum b-lactamase–producing Klebsiella pneumoniae in mice. Antimicrob Agents Chemother 2005; 49:3513–6. 42. Pultz NJ, Donskey CJ. Effect of antibiotic treatment on growth of and toxin production by Clostridium difficile in the cecal contents of mice. Antimicrob Agents Chemother 2005; 49:3529–32. 43. Hentges DJ, Stein AJ, Casey SW, Que JU. Protective role of intestinal flora against infection with Pseudomonas aeruginosa in mice: influence of antibiotics on colonization resistance. Infect Immun 1985; 47:118–22. 44. Sullivan A, Edlund C, Nord CE. Effect of antimicrobial agents on the ecological balance of human microbiota. Lancet Infect Dis 2001; 1: 101–14. 45. Leibovitz A, Dan M, Zinger J, Carmeli Y, Habot B, Segal R. Pseudomonas aeruginosa and oropharyngeal ecosystem of tube-fed patients. Emerg Infect Dis 2003; 9:956–9. 46. Simon GL, Gorbach SL. Normal alimentary tract microflora. In: Blaser MJ, Smith PD, Ravdin JI, Greenberg HB, Guerrant RL, eds. Infections of the gastrointestinal tract. 2nd ed. New York: Raven Press; 1995: 53–69. 47. Edlund C, Nord CE. Effect of quinolones on intestinal ecology. Drugs 1999; 58(Suppl 2):65–70. 48. Joris JJ, Van de Leur PM, Vollaard EJ, Janssen AJHM, Dofferhoff ASM. Influence of low dose ciprofloxacin on microbial colonization of the digestive tract in healthy volunteers during normal and during impaired colonization resistance. Scand J Infect Dis 1997; 29:297–300. 49. Bhalla A, Pultz NJ, Ray AJ, Hoyen CK, Eckstein EC, Donskey CJ. Antianaerobic antibiotic therapy promotes overgrowth of antibioticresistant gram-negative bacilli and vancomycin-resistant enterococci in the stool of colonized patients. Infect Control Hosp Epidemiol 2003; 24:644–9. 50. Boyce JM, Pitet D, Healthcare Infection Control Practices Advisory Committee, HICPAC/SHEA/APIC/IDSA Hand Hygiene Task Force. Guideline for hand hygiene in health-care settings. Recommendations of the Healthcare Infection Control Practices Advisory Committee and the HIPAC/SHEA/APIC/IDSA Hand Hygiene Task Force. Am J Infect Control 2002; 30:S1–46. 51. Wright MO, Hebden JN, Harris AD, et al. Aggressive control measures for resistant Acinetobacter baumannii and the impact on acquisition of methicillin-resistant Staphylococcus aureus and vancomycin-resistant Enterococcus in a medical intensive care unit. Infect Control Hosp Epidemiol 2004; 25:167–8. 52. Lemmen SW, Hafner H, Zolldann D, Stanzel S, Lutticken R. Distribution of multi-resistant gram-negative versus gram-positive bacteria in the hospital inanimate environment. J Hosp Infect 2004; 56:191–7. 53. Denton M, Wilcox MH, Parnell P, et al. Role of environmental cleaning in controlling an outbreak of Acinetobacter baumannii on a neurosurgical intensive care unit. J Hosp Infect 2004; 56:106–10. 54. Reuter S, Sigge A, Wiedeck H, Trautmann M. Analysis of transmission pathways of Pseudomonas aeruginosa between patients and tap water outlets. Crit Care Med 2002; 30:2222–8. 55. Wang SH, Sheng WH, Chang YY, et al. Healthcare-associated outbreak due to pan–drug resistant Acinetobacter baumannii in a surgical intensive care unit. J Hosp Infect 2003; 53:97–102. Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 19. Quinn JP, DiVincenzo CA, Foster J. Emergence of resistance to ceftazidime during therapy for Enterobacter cloacae infections. J Infect Dis 1987; 155:942–7. 20. Quinn JP, Miyashiro D, Sahm D, Flamm R, Bush K. Novel plasmidmediated b-lactamase (TEM-10) conferring selective resistance to ceftazidime and aztreonam in clinical isolates of Klebsiella pneumoniae. Antimicrob Agents Chemother 1989; 33:1451–6. 21. Quinn JP, Dudek EF, DiVincenzo CA, Lucks DA, Lerner SA. Emergence of resistance to imipenem during therapy for Pseudomonas aeruginosa infections. J Infect Dis 1986; 154:289–94. 22. Ahmad M, Urban C, Mariano N, et al. Clinical characteristics and molecular epidemiology associated with imipenem-resistant Klebsiella pneumoniae. Clin Infect Dis 1999; 29:352–5. 23. Corbella X, Montero A, Pujol M, et al. Emergence and rapid spread of carbapenem resistance during a large and sustained hospital outbreak of multiresistant Acinetobacter baumannii. J Clin Microbiol 2000; 38: 4086–95. 24. Chow JW, Fine MJ, Shlaes DM, et al. Enterobacter bacteremia: clinical features and emergence of antibiotic resistance during therapy. Ann Intern Med 1991; 115:585–90. 25. Prevot M, Andremont A, Sancho-Garnier H, Tancrede C. Epidemiology of intestinal colonization by members of the family Enterobacteriaceae resistant to cefotaxime in a hematology-oncology unit. Antimicrob Agents Chemother 1986; 30:945–7. 26. Richard P, Delangle MH, Raffi F, Espaze E, Richet H. Impact of fluoroquinolone administration on the emergence of fluoroquinoloneresistant gram-negative bacilli from gastrointestinal flora. Clin Infect Dis 2001; 32:162–6. 27. Landman D, Bratu S, Alam M, Quale J. Citywide emergence of Pseudomonas aeruginosa strains with reduced susceptibility to polymyxin B. J Antimicrob Chemother 2005; 55:954–7. 28. Harbarth S, Harris AD, Carmeli Y, Samore MH. Parallel analysis of individual and aggregated data on antibiotic exposure and resistance in gram-negative bacilli. Clin Infect Dis 2001; 33:1462–8. 29. Donskey CJ, Chowdhry TK, Hecker MT, et al. Effect of antibiotic therapy on the density of vancomycin-resistant enterococci in the stool of colonized patients. N Engl J Med 2000; 343:1925–32. 30. D’Agata EM, Gautam S, Green WK, Tang YW. High rate of falsenegative results of the rectal swab culture method in detection of gastrointestinal colonization with vancomycin-resistant enterococci. Clin Infect Dis 2002; 34:167–72. 31. Asensio A, Oliver A, Gonzalez-Diego P, et al. Outbreak of a multiresistant Klebsiella pneumoniae strain in an intensive care unit: antibiotic use as a risk factor for colonization and infection. Clin Infect Dis 2000; 30:55–60. 32. Harris AD, Perencevich E, Roghmann M, Morris G, Kaye KS, Johnson JA. Risk factors for piperacillin-tazobactam–resistant Pseudomonas aeruginosa among hospitalized patients. Antimicrob Agents Chemother 2002; 46:854–8. 33. Kaye KS, Harris AD, Gold H, Carmelli Y. Risk factors for recovery of ampicillin-sulbactam–resistant Escherichia in hospitalized patients. Antimicrob Agents Chemother 2000; 44:1004–9. 34. Schwaber MJ, Graham CS, Sands BE, Gold HS, Carmelli Y. Treatment with a broad-spectrum cephalosporin versus piperacillin-tazobactam and the risk for isolation of broad-spectrum cephalosporin–resistant Enterobacter species. Antimicrob Agents Chemother 2003; 47:1882–6. 35. Bonten MJ, Bergmans DC, Speijer H, Stobberingh EE. Characteristics of polyclonal endemicity of Pseudomonas aeruginosa colonization in intensive care units. Am J Respir Crit Care Med 1999; 160:1212–9. 36. Paterson DL, Singh N, Rihs JD, Squier C, Rihs BL, Muder RR. Control of an outbreak of infection due to extended-spectrum b-lactamase–producing Escherichia coli in a liver transplant unit. Clin Infect Dis 2001; 33:126–8. 37. Paterson DL, Mulazimoglu L, Casellas JM, et al. Epidemiology of ciprofloxacin resistance and its relationship to extended-spectrum b-lactamase production in Klebsiella pneumoniae isolates causing bacteremia. Clin Infect Dis 2000; 30:473–8. 63. Hecker MT, Aron DC, Patel NP, Lehman MK, Donskey CJ. Unnecessary use of antimicrobials in hospitalized patients: current patterns of misuse with an emphasis on the antianaerobic spectrum of activity. Arch Intern Med 2003; 163:972–8. 64. Chastre J, Wolff M, Fagon JY, et al. Comparison of 8 vs 15 days of antibiotic therapy for ventilator-associated pneumonia in adults: a randomized trial. JAMA 2003; 290:2588–98. 65. Paterson DL, Yu VL. Editorial response: extended-spectrum b-lactamases: a call for improved detection and control. Clin Infect Dis 1999; 29:1419–22. 66. Aubert G, Carricajo A, Vautrin AC, et al. Impact of restricting fluoroquinolone prescription on bacterial resistance in an intensive care unit. J Hosp Infect 2005; 59:83–9. 67. Rahal JJ, Urban C, Horn D, et al. Class restriction of cephalosporin use to control total cephalosporin resistance in nosocomial Klebsiella. JAMA 1998; 280:1233–7. 68. Harris AD, Bradham DD, Baumgarten M, et al. The use and interpretation of quasi-experimental studies in infectious diseases. Clin Infect Dis 2004; 38:1586–91. 69. Stiefel U, Pultz NJ, Harmoinen J, Koski P, Lindevall K, Donskey CJ. Oral administration of recombinant b-lactamase prevents piperacillininduced overgrowth of nosocomial pathogens in the intestinal tract of mice. J Infect Dis 2003; 188:1605–9. 70. Wilson M. Light-activated antimicrobial coating for the continuous disinfection of surfaces. Infect Control Hosp Epidemiol 2003; 24:782–4. Antibiotics and Gram-Negative Bacilli • CID 2006:43 (Suppl 2) • S69 Downloaded from http://cid.oxfordjournals.org/ by guest on September 9, 2014 56. Pimentel JD, Low J, Styles K, Harris OC, Hughes A, Athan E. Control of an outbreak of multi-drug–resistant Acinetobacter baumannii in an intensive care unit and a surgical ward. J Hosp Infect 2005; 59:249–53. 57. Koeleman JG, Parlevliet GA, Dijkshoorn L, Savelkoul PH, Vandenbroucke-Grauls CM. Nosocomial outbreak of multi-resistant Acinetobacter baumannii on a surgical ward: epidemiology and risk factors for acquisition. J Hosp Infect 1997; 37:113–23. 58. Urban C, Segal-Maurer S, Rahal JJ. Considerations in control and treatment of nosocomial infections due to multidrug-resistant Acinetobacter baumannii. Clin Infect Dis 2003; 36:1268–74. 59. Borer A, Gilad J, Megreleshvili R, et al. Prevalence and control of Acinetobacter baumannii skin colonization among medical intensivecare unit patients [abstract K-1100]. In: Program and abstracts of the 43rd Interscience Conference on Antimicrobial Agents and Chemotherapy. Washington, DC: American Society for Microbiology, 2003. 60. De Jonge E, Schultz MJ, Spanjaard L, et al. Effects of selective decontamination of digestive tract on mortality and acquisition of resistant bacteria in intensive care: a randomized controlled trial. Lancet 2003; 362:1011–6. 61. Jackson RJ, Smith SD, Rowe MI. Selective bowel decontamination results in gram-positive translocation. J Surg Res 1990; 48:444–7. 62. Bergmans DC, Bonten MJ, Gaillard CA, et al. Prevention of ventilatorassociated pneumonia by oral decontamination: a prospective, randomized, double-blind, placebo-controlled study. Am J Respir Crit Care Med 2001; 164:382–8.
© Copyright 2018