Antibiotic Regimens and Intestinal Colonization with Antibiotic-Resistant Gram-Negative Bacilli

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 [9]. Finally, the intestinal tract provides an important site for transfer of genes
conferring antibiotic resistance [10].
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 (curtisd123@yahoo.com).
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
[11]. Escherichia coli, a member of the indigenous colonic microflora, is normally maintained at relatively
low population densities by the predominant anaerobic
microflora [12]. 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 [13]. Foods such as salads may contain relatively large numbers of P. aeruginosa or Entero-
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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) [14], 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
[16]. 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) [17].
ANTIBIOTICS AND COLONIZATION
WITH GRAM-NEGATIVE BACILLI
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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 [18]. 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 [9]. 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 [9].
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 [7].
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 [7].
Similarly, piperacillin-tazobactam, imipenem, aminoglycosides,
vancomycin, and broad-spectrum cephalosporins have all been
associated with piperacillin- and tazobactam-resistant P. aeruginosa [32].
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. [33] 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. [34]. Paterson et
al. [36] 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 [7].
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. [43]
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
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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 [44]. 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 [40], 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 [44]. 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 [44]. 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 [45]. Second, patients often receive proton pump inhibitors or histamine2 blockers that inhibit production of stomach
acid [9]. These agents promote overgrowth of gram-negative
bacilli in the stomach and facilitate passage of organisms into
the small intestine [9]. Third, the colonic microflora of ill hospitalized patients may be altered in the absence of antibiotic
therapy [46]. 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. [48].
Effect of antibiotics with activity against intestinal anaerobes. Antibiotics that inhibit intestinal anaerobes promote
overgrowth of VRE in stools of colonized patients [29]. 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 [49]. 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 [49].
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 [50].
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 [50]. For organisms resistant to multiple
antibiotics or during outbreaks, contact precautions may be indicated [9]. 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. [6] 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 [9], 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
[9]. A recent study found that patients harboring antibioticAntibiotics and Gram-Negative Bacilli • CID 2006:43 (Suppl 2) • S65
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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. [48] 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 [48].
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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 [49]).
resistant gram-negative pathogens were less likely to have environmental contamination than were those harboring resistant
gram-positive pathogens (4.9% vs. 24.7%) [52]. However, extensive environmental contamination with Acinetobacter baumannii has been described, and this organism survives for prolonged periods on surfaces [53]. In addition, P. aeruginosa may
persistently contaminate moist areas, such as sinks [54]. 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. [59] 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. [58] 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
[9]. 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 [60]. 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) [60]. Jackson et al. [61] 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 [36]. 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 [62].
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 [63]. 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 [69]. Others have developed light-activated antimicrobial coatings for continuous disinfection of surfaces [70].
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. [64] 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 [65]. 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 [66]. Unfortunately, substitution of one agent
for another may lead to the emergence of resistance to the new
agent [67]. 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 [68].
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.