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Journal of Clinical Microbiology, December 2003, p. 5530-5536, Vol. 41, No. 12
0095-1137/03/$08.00+0     DOI: 10.1128/JCM.41.12.5530-5536.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

Nosocomial Outbreak of Infections by Proteus mirabilis That Produces Extended-Spectrum CTX-M-2 Type ß-Lactamase

Noriyuki Nagano,^1* Naohiro Shibata,² Yuko Saitou,¹ Yukiko Nagano,¹ and Yoshichika Arakawa²

Medical Microbiology Laboratory, Funabashi Medical Center, 1-21-1 Kanasugi, Funabashi, Chiba,¹ Department of Bacterial Pathogenesis and Infection Control, National Institute of Infectious Diseases, Tokyo, Japan²

Received 9 May 2003/ Returned for modification 25 August 2003/ Accepted 18 September 2003

Top Abstract Introduction Materials and Methods Results Discussion References

 
Nineteen multidrug-resistant Proteus mirabilis strains were isolated from 19 patients suffering from infections probably caused by P. mirabilis. These strains were recovered from urine or other urogenital specimens of 16 inpatients and three outpatients with a hospitalization history in a urology ward of Funabashi Medical Center, from July 2001 to August 2002. These strains demonstrated resistance to cefotaxime, ceftriaxone, cefpodoxime, and aztreonam, while they were highly susceptible to ceftazidime (MIC, 0.5 µg/ml). The resistance level of these strains to cefotaxime was decreased by the presence of clavulanic acid. Therefore, the strains were speculated to produce extended-spectrum class A ß-lactamases. These strains were later found to carry bla_CTX-M-2 genes by both PCR and sequencing analyses. The profiles of SmaI-digested genomic DNA of 19 isolates were distinguished into five different clusters by biased sinusoidal field gel electrophoresis. Four of them, consisting of 18 isolates, were suggested to be a clonal expansion. These findings suggested that a nosocomial outbreak of infections by CTX-M-2-producing P. mirabilis had occurred in our medical center. Most patients suffered from urogenital malignancies with long-term catheterization. Cefazolin, cefoperazone-sulbactam, and/or levofloxacin were mostly administered to the patients, but these agents seemed ineffective for eradication of CTX-M-2 producers. Early recognition and rapid identification of colonizing antimicrobial-resistant bacteria, including CTX-M-2-producing P. mirabilis, would be the most effective measures to cope with further spread of this kind of hazardous microorganism in clinical environments.

Top Abstract Introduction Materials and Methods Results Discussion References

 
The increasing prevalence of plasmid-mediated extended-spectrum ß-lactamases (ESBLs) in members of the family Enterobacteriaceae has become a serious clinical problem on a worldwide scale (8). ESBLs of Ambler's molecular class A (1) belonging to Bush's functional group 2be (10) are capable of hydrolyzing a wide range of ß-lactams, including oxyimino-ß-lactams and monobactam, but usually remain ineffective against cephamycins such as cefoxitin, cefmetazole, and cefotetan as well as carbapenems. These class A ß-lactamases tend to be blocked by ß-lactamase inhibitors such as clavulanic acid (10). The majority of ESBLs are derivatives of TEM-1, TEM-2, or SHV-1 enzymes, resulting from a few amino acid substitutions (10). In contrast to these TEM- and SHV-derived ESBLs, CTX-M type ß-lactamases, which constitute a new family of class A enzymes, are exclusively active against cefotaxime compared to other oxyimino-cephalosporins, including ceftazidime (39).

More than 30 CTX-M-type ß-lactamases have so far been described in various species of Enterobacteriaceae but mostly in Salmonella enterica serovar Typhimurium, Escherichia coli, and Klebsiella pneumoniae since the initial reports of Toho-1-producing E. coli in Japan (18) and CTX-M-1/MEN-1 in 1989 in Germany and France (2, 3). Strains producing other CTX-M enzymes have been isolated in separate geographic areas, including Europe (9, 15, 16, 35, 40), South America (4, 7, 30), and the Middle and Far East (4, 18, 23, 41). CTX-M-type ß-lactamases can be classified into four clusters according to their amino acid sequences: CTX-M-1-group, with CTX-M-1 (4), -M-3 (16), -M-10 (27), -M-11 (GenBank accession no. AY005110), -M-12 (21), -M-15 (20), -M-22 (GenBank accession no. AY080894), and -M-23 (GenBank accession no. AF488377); CTX-M-2-group, with CTX-M-2 (4), -M-4 (15), -M-5 (9), -M-6 (14), -M-7 (14), -M-20 (36), -M-24 (GenBank accession no. AY143430), and Toho-1 (18); CTX-M-8 group (7); and CTX-M-9 group, with CTX-M-9 (35), -M-13 (12), -M-14/18 (29), -M-16 (6), -M-19 (32), -M-21 (36), and Toho-2 (23).

In this paper, we report a nosocomial outbreak of infections caused by CTX-M-2 ß-lactamase-producing Proteus mirabilis in a urology ward. P. mirabilis is one of the most common causes of urinary tract infections. Because of the difficulty in eradicating P. mirabilis species from immunocompromised hosts (13), this bacterial species is usually considered an important cause of nosocomial infections (34). Although the most predominant plasmid-mediated ß-lactamases found in clinical isolates of P. mirabilis are TEM-derived ESBLs (5, 11, 22, 24, 28, 33), the emergence of CTX-M-type enzymes with extended substrate specificity has been a serious concern (7, 36). In the present study, we investigated the CTX-M-2-producing P. mirabilis strains that caused a nosocomial outbreak in a urology ward in our medical center.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Patients and bacterial strains. From July 2001 to August 2002, 19 nonduplicated multiresistant P. mirabilis clinical strains were isolated from 19 patients suffering from infections probably caused by P. mirabilis. These strains were recovered from urine or other urogenital specimens of 16 inpatients and three outpatients with a hospitalization history in a urology ward of Funabashi Medical Center. This hospital has 426 beds and serves as an acute-care municipal hospital for a population of 560,000 in Funabashi City, Chiba, Japan. Tables 1 and 2 show the clinical background of patients for each isolate and their respective treatment outcomes. All 19 isolates were suggested to produce inhibitor-susceptible class A ß-lactamase based on the double-disk synergy test results. Biochemical identification of isolates was performed with an NEG Combo 5J panel and WalkAway-96 SI System (Dade Behring, Sacramento, Calif.) according to the manufacturer's instructions. ß-Lactamase testing was performed based on microacidimetry with a commercial product (P/Case Test; Nissui Pharmaceutical, Tokyo, Japan). Bacterial strains were stored before use in Casitone medium (Eiken Chemical, Tokyo, Japan) at room temperature.

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TABLE 1. Origins of CTX-M-2-producing P. mirabilis isolates and medical records of patients

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TABLE 2. Treatment outcome for patients with respect to detection of CTX-M-2-producing P. mirabilis isolates

Antimicrobial susceptibility testing. MICs were determined by a microdilution broth method with a WalkAway-96 SI System (NEG Combo 5J and NEG MIC 5J panels; Dade Behring) with an inoculum of 10⁴ CFU per well. Susceptibility categories were determined according to the National Committee for Clinical Laboratory Standards (NCCLS) criteria (26).

ESBL plus Panel (Dade Behring) with an inoculum of 10⁴ CFU per well was used complementarily for MIC measurements, with incubation for 18 h at 35°C, and then assessed visually.

Double-disk synergy test. For screening ESBL-producing strains, the double-disk synergy test was used. Antimicrobial disks for Mueller-Hinton agar (BBL Microbiology Systems, Cockeysville, Md.) tests, cefotaxime (30 µg), ceftazidime (30 µg), aztreonam (30 µg), and amoxicillin-clavulanic acid (20 µg and 10 µg) were obtained from Nissui Pharmaceutical. The distance between disks was adjusted so that synergy could be detected accurately (38).

PCR analysis. A search for bla_TEM, bla_SHV, bla_CTX-M-1, bla_CTX-M-2, and bla_CTX-M-9 genes in clinical isolates was performed by PCR amplification with the following sets of primers: 5'-CCGTGTCGCCCTTATTCC-3' and 5'-AGGCACCTATCTCAGCGA-3' for bla_TEM, 5'-ATTTGTCGCTTCTTTACTCGC-3' and 5'-TTTATGGCGTTACCTTTGACC-3' for bla_SHV, 5'-CGGTGCTGAAGAAAAGTG-3' and 5'-TACCCAGCGTCAGATTAC-3' for bla_CTX-M-1, 5'-ACGCTACCCCTGCTATTT-3' and 5'-CCTTTCCGCCTTCTGCTC-3' and for bla_CTX-M-2, and 5'-GCAGATAATACGCAGGTG-3' and 5'-CGCCGTGGTGGTGTCTCT-3' for bla_CTX-M-9. Freshly isolated colonies were suspended in distilled water and adjusted to a 0.5 MacFarland, which was boiled for 10 min. Supernatant obtained after centrifugation at 13,000 rpm for 5 min was used as template DNA.

PCRs were carried out in 50-µl volumes containing 5 µl of DNA, 0.5 µM each primer, 200 µM deoxynucleoside triphosphates, 1.25 U of TaKaRa Ex Taq (Takara), and PCR buffer (Takara) with the following parameters: initial denaturation at 94°C for 2 min, denaturation at 94°C for 1 min, primer annealing at 55°C for 1 min, and extension at 72°C for 1.5 min, repeated for 30 cycles; and a final extension at 72°C for 5 min.

CTX-M-2-specific PCR and DNA sequencing. Amplification of the bla_CTX-M-2 gene and flanking regions was carried out with the oligonucleotide primers M-2-F (5'-TTCGCCGCTCAATGTTA-3') and M-2-R (5'-GCATCAGAAACCGTGGG-3'), corresponding to nucleotides 22 to 38 and 852 to 868, respectively, of the structural gene. Plasmid DNA was prepared from each isolate by the Kado and Liu method (19) and used as templates for PCR analyses. PCRs were performed as described above. Cycling conditions were denaturation at 94°C for 1 min, annealing at 55°C for 1 min, and extension at 72°C for 1.5 min, repeated for 30 cycles. PCR-generated amplicons were purified with a QIAquick PCR Purification Kit (Qiagen Inc., Valencia, Calif.), and sequenced directly on both strands with a BigDye terminator cycle sequencing ready reaction kit and ABI 3100 DNA sequencer (Applied Biosystems, Foster City, Calif.).

Genomic typing. Chromosomal DNAs from clinical isolates embedded in agarose gel plugs (InCert; Bio-Whittaker Molecular Applications, Rockland, Maine) were subjected to treatments with lysozyme and sodium dodecyl sulfate containing proteinase K, then incubated overnight at 30°C with 12.5 U of SmaI (Takara Shuzo Co., Kyoto, Japan). Plugs were mounted into the wells of a 1% SeaKem GTG Agarose (Bio-Whittaker) in 50 mM Tris-borate-EDTA buffer (pH 8.4). The biased sinusoidal field gel electrophoresis system (Atto Corp., Tokyo, Japan) (25), a modified pulsed-field gel electrophoresis technique utilizing a biased sinusoidal electric field for separation of large DNA molecules, was employed at 12°C with the field parameters Eb = 1.2 V/cm and Es = 7.3 V/cm. Lambda DNA ladders (48.5 kb to 1 Mb; Takara) were used as molecular size markers.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Bacterial strains and clinical features. Multiresistant P. mirabilis isolates were obtained from 16 inpatients in a urology ward and three outpatients with a hospitalization history in that ward, 44 to 83 years old, 16 males and 3 females, from July 2001 to August 2002 (Table 1). Of 19 nonduplicated isolates, 9 isolates (strains 2, 3, 5, 6, 7, 13, 14, 17, and 19) were recovered from indwelling catheter urine samples, six isolates (strains 4, 8, 9, 11, 12, and 18) were from midstream urine samples, two isolates (strains 10 and 16) from catheter urine samples, one (strain 1) from an operative wound, and one (strain 15) from sputum. The common underlying disease was urogenital malignancies, including bladder cancer in nine patients, prostatic cancer in four patients, followed in order by renal failure in three, bladder stone in two, and stomach cancer as well as pyelonephritis in one patient. Cefazolin and cefoperazone-sulbactam had been administered most frequently in 12 patients, and levofloxacin had been prescribed to 11 patients within 30 days before isolation of the multiresistant P. mirabilis.

Table 2 shows the outcomes of antibiotic therapy for 19 patients after detection of the isolates, in which 12 patients were traceable for evaluation. Eradication was achieved in four patients with sulfamethoxazole-trimethoprim and flomoxef for patient 7, flomoxef and levofloxacin for patient 12, cefoperazone-sulbactam and ceftazidime for patient 13, and kanamycin, metronidazole, and levofloxacin for patient 19. In eight patients, P. mirabilis isolates were persistently detected despite the therapy with cefoperazone-sulbactam (patients 1, 4, 5, and 9), flomoxef (patient 1), imipenem/cilastatin (patient 5), and ceftazidime (patient 14). Treatment outcome could not be evaluated in seven patients due to lack of bacteriological follow-up in four and death from underlying disease in three patients. In 11 of 19 patients, other bacterial species were isolated besides P. mirabilis; Enterococcus faecalis, methicillin-resistant Staphylococcus aureus, and Pseudomonas aeruginosa were predominant. Moreover, no ESBL producers were detected among eight bacterial strains belonging to the family Enterobacteriaceae or P. aeruginosa isolated from 19 patients.

Determination of antibiotic susceptibility. MICs of antimicrobial agents for 19 clinical isolates are shown in Table 3. These isolates showed very similar susceptibility profiles, characterized by elevated MICs of cefotaxime (MICs, >128 µg/ml), ceftriaxone (MICs, >64 µg/ml), cefpodoxime (MICs, >64 µg/ml), aztreonam (MICs, 8 to >16 µg/ml), while they were susceptible to ceftazidime (MICs, 0.5 µg/ml). For all isolates, the MICs of cefotaxime were decreased drastically to 0.12 µg/ml in the presence of 4 µg of clavulanic acid per ml, whereas the MICs of ceftazidime for these strains were not obviously influenced by the presence of clavulanic acid. The MICs of cefoperazone-sulbactam were 8 to 32 µg/ml. MICs of other ß-lactams, imipenem and melopenem, for the same strains were 2 µg/ml and 0.5 µg/ml, respectively. There was a trend towards resistance to gentamicin (MICs, 2 to >8 µg/ml), minocycline (MICs, >8 µg/ml), and levofloxacin (MICs, 2 to >4 µg/ml) among the isolates.

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TABLE 3. Antibiotic susceptibilities of clinical P. mirabilis isolates presented in

ß-Lactamase study. The production of ß-lactamase was detected by the P/Case test, which can distinguish between penicillinase (benzylpenicillin as the substrate) and cephalosporinase (cephaloridine and clavulanic acid as the substrate). Penicillinase production was detected in all 19 strains tested. With the double-disk synergy test, expanded growth-inhibitory zones indicative of class A ß-lactamase production were observed with cefotaxime, ceftazidime, and aztreonam disks among all 19 strains (data not shown). MICs of cefotaxime (>128 µg/ml) decreased dramatically to 0.12 µg/ml in the presence of 4 µg of clavulanic acid per ml, suggesting the production of a CTX-M type class A ß-lactamase (Table 2).

PCR and sequencing of bla_CTX-M-2 gene. The preliminary PCR search revealed that all 19 P. mirabilis isolates showed 780-bp amplification products for bla_CTX-M-2 genes, whereas no amplicons were observed for bla_TEM, bla_SHV, bla_CTX-M-1, or bla_CTX-M-9. The entire coding sequences of the bla_CTX-M-2 gene and flanking regions were subsequently amplified with more specific primers for bla_CTX-M-2, M-2-F and M-2-R, and sequenced on both strands. The BLAST analysis of the nucleotide sequences and the deduced protein sequences showed that P. mirabilis isolates produced CTX-M-2 group ß-lactamase (4).

SmaI-digested genomic DNA profiles. The SmaI-digested genomic DNAs of 19 clinical isolates were classified into five different clusters (Fig. 1). Thirteen isolates of strains 1, 3, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, and 17 had the same restriction profiles (pattern I), while the patterns of strains 15, 18, and 19 differed from pattern I in only one band (pattern II). The patterns of strains 4 (pattern III) and 16 (pattern IV) were different from each other in three bands and differed in two bands from pattern I. Therefore, these 18 isolates were clonally related. The pattern of strain 2 (pattern V) was completely distinguishable from those of 18 isolates (Fig. 1).

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FIG. 1. Profiles of the genomic DNAs of 19 P. mirabilis isolates producing CTX-M-2 ß-lactamase after digestion with SmaI. See Table 1 for the origins and backgrounds of the isolates. Lane M, lambda DNA ladder as molecular size markers.

Top Abstract Introduction Materials and Methods Results Discussion References

 
ESBL production in P. mirabilis was first documented in 1993 (17), and the increase in clinical prevalence of ESBL-producing strains has recently been noted in survey studies in separate geographic areas, including the United States, Europe, and Asia. The proportion of ESBL-positive isolates has increased from 0.8% of P. mirabilis isolates in 1991 (17) to 6.9% in 1998 (13) in France. Surveillance studies conducted in the United States and Italy showed 9.5% and 8.8% ESBL prevalence among P. mirabilis isolates, respectively (22, 37). TEM-derived ESBLs showing a wide diversity were the most predominant among ESBLs (5, 11, 22, 24, 28, 33), but other enzymes belonging to group 2be (7, 36) have also been observed in P. mirabilis. Because of the production of such diverse class A ß-lactamases in P. mirabilis, as well as its predilection for the urinary tract, the emergence and proliferation of multidrug resistant P. mirabilis could pose a threat, especially in catheterized patients with malignancy as a cause of subsequent nosocomial infections.

To our knowledge, this is the first report of a nosocomial outbreak of infections caused by CTX-M-2-producing P. mirabilis strains in a Japanese medical institution. Nineteen isolates found in its urology ward were initially speculated to produce CTX-M-type class A ß-lactamase, since consistent high MICs of cefotaxime, ceftriaxone, cefpodoxime, and aztreonam were observed for these strains. The drastic reduction in the MICs of cefotaxime in the presence of clavulanic acid supported this speculation. Guidelines for screening and confirmatory tests for ESBL producers have been established by the National Committee for Clinical Laboratory Standards (26). These guidelines apply specifically to K. pneumoniae, E. coli, and Klebsiella oxytoca. However, the incidence of ESBLs producers has also been increasing in many other genera belonging to the family Enterobacteriaceae such as Citrobacter, Enterobacter, Morganella, Proteus, Providencia, Salmonella, Serratia, and other gram-negative bacilli (8). This is also the case in our medical center, where nosocomial infection due to ESBL-producing Acinetobacter baumannii has recently been identified (unpublished data).

In our experience, the NCCLS guidelines could be applicable for detection of CTX-M-2 producers among P. mirabilis that produce no intrinsic AmpC cephalosporinases, although some modification in the NCCLS criteria might be needed. This would be of critical importance to be able to detect ESBL and CTX-M-type ß-lactamase producers for effective clinical management of patients with infections by reliable and appropriate therapeutic options. Thus, accurate monitoring of ESBL prevalence would be mandatory to promote hospital infection control procedures.

The production of ESBL is alternatively confirmed by the double-disk synergy test, by which a synergistic effect on growth inhibition is observed with the coexistence of clavulanic acid and broad-spectrum cephalosporins, including cefotaxime, ceftazidime, or aztreonam. This test was also useful to detect CTX-M-2 production in P. mirabilis strains. The NCCLS recommends, however, reporting that Klebsiella spp. and E. coli strains producing ESBLs may be clinically resistant to therapy with penicillins, cephalosporins, or aztreonam, despite apparent in vitro susceptibility to some of these agents. This recommendation might cause confusion, implying that ceftazidime may be ineffective for treatment of infections caused by cefotaxime-resistant strains that produce CTX-M-type ß-lactamase despite the fact that strains producing only CTX-M-type ß-lactamase seem highly susceptible to ceftazidime.

As a practical matter, eradication of CTX-M-2-producing P. mirabilis isolates was successfully achieved by therapy with ceftazidime in patient 13, although combination therapy with cefoperazone-sulbactam was employed in this case. However, ceftazidime therapy alone failed to eradicate the organism in patient 14 (Table 2). There was a difference in treatment regimens between these two cases. Continuous administration of ceftazidime (1 g/day intravenously) for 8 days was used for patient 13, while two courses of 3-day-repeated administration of the same dosage at a 19-day interval was administered to patient 14. Furthermore, while patient 13 was infected only with P. mirabilis, patient 14 had polymicrobial infection with E. faecalis, Staphylococcus haemolyticus, Enterobacter cloacae, and methicillin-resistant S. aureus in addition to CTX-M-2-producing P. mirabilis. This, together with the incomplete antibiotic therapy, might explain the poor therapeutic response to ceftazidime in patient 14. To our knowledge, no clinical evidence obtained by double-blind clinical trials supporting the ineffectiveness of ceftazidime for infections with CTX-M-type producers has been reported. Clinical studies to address this issue should be conducted immediately to either corroborate or call into question the NCCLS recommendation.

Most patients from whom CTX-M-2-producing P. mirabilis strains were isolated had urological malignancies and long-term catheterization. All patients had received antibiotic therapy for 30 days before isolation of P. mirabilis, in which cefazolin, cefoperazone-sulbactam, and/or levofloxacin had been most frequently administered. Among the antibiotics used after detection of the organisms, cefoperazone-sulbactam, imipenem/cilastatin, and flomoxef showed low MICs. However, eradication with cefoperazone-sulbactam was noted in only one (patient 13) of five patients, and it was ineffective in combination with isepamicin. Since CTX-M-type enzymes can hydrolyze cefoperazone and tend to be hardly blocked by sulbactam, CTX-M producers usually demonstrate insusceptibility or resistance to the combination of cefoperazone-sulbactam (41). Thus, random or uniform prescription of cefoperazone-sulbactam may induce nosocomial spread of gram-negative bacilli which produce CTX-M-type enzymes.

Actually, 12 of 19 patients were prescribed cefoperazone-sulbactam prior to the isolation of CTX-M-2-producing P. mirabilis. The imipenem/cilastatin used in one patient (patient 5) proved in ineffective. Flomoxef was used in three patients (patients 1, 7, and 12), and eradication was achieved in the latter two cases. Levofloxacin, which tends to be preferentially used in urinary tract infections in Japan, showed high MICs for P. mirabilis isolates and seemed ineffective for most of the eight patients for whom it was used. These findings suggest difficulties in eradication of CTX-M-2-producing P. mirabilis strains even with antibiotics with low MICs. The biofilm-forming ability of bacteria, including P. mirabilis, on urinary catheters may be one reason for the failure to eradicate the organisms.

The restriction profiles of genomic DNAs from 19 P. mirabilis isolates shared concordant patterns (pattern I) among 13 isolates (strains 1, 3, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, and 17), suggesting a clonal expansion of CTX-M-2 producers. Moreover, a clonal relatedness among five isolates (strains 15, 18, 19, 4, and 16 corresponding to patterns II, III, and IV) was also indicated. These five strains presumably relate to the previous 13 strains with pattern I. However, the pattern of strain number 2 (pattern V) was apparently distinct from those of the other 18 isolates classified as patterns I, II, III, and IV. These findings suggested that a nosocomial outbreak of 18 clonally related isolates and one isolate subjected to a different genetic lineage had occurred in a certain ward carrying the same CTX-M-2 ß-lactamase determinant.

The Dienes test, which visualizes a unique feature of the swarming ability of P. mirabilis, has been utilized for an epidemiologic typing method (31), which we adopted in the present study for initial epidemiologic characterization of isolates. The 16 isolates demonstrating pattern I or II were indistinguishable and formed one Dienes type, whereas one isolate with patterns III and one isolate with pattern V formed independent Dienes type. Furthermore, one isolate subjected to pattern IV showed less than detectable swarming ability (data not shown). While the Dienes test was indeed partially applicable for typing of P. mirabilis, the biased sinusoidal field gel electrophoresis employed in this study was much more useful for the epidemiological analyses, although it requires skill and involves somewhat complicated procedures.

Infection control at the initial stage of the outbreak was difficult due to frequent patient transfers within the urology ward. Eventually, the outbreak was successfully brought under control by intensive surveillance, improvement of facilities including disinfection equipment, prudent use of antibiotics, and due precautions to prevent contact transmission of microorganisms. The most effective measures to prevent the further spread of CTX-M-2 ß-lactamase-producing P. mirabilis were rapid identification of colonization status of such bacteria among all immunocompromised patients with severe urological disorders by periodic urine culturing at admission and once-per-week follow-up testing with informed consent.

In our medical center, first- and third-generation cephalosporins have been preferentially used as first-line drugs. Carbapenems and penicillins as well as first- and third-generation cephalosporins have accounted for the great majority. These antibiotics might well provide selective pressure for proliferation of the CTX-M-2 producers. Early recognition of bacterial strains possessing antimicrobial resistance would contribute to an appropriate antibiotic treatment regimen that would be essential for prevention of nosocomial infections.

 
We are grateful to N. Sato and Y. Watanabe, Department of Urology; Y. Suzuki, H. Tabeta, H. Kamii, M. Shoji, and K. Iwata, Infection Control Team; and N. Suwa, and M. Toyama, Medical Microbiology Laboratory, Funabashi Medical Center, for their careful infection control.

This work was supported in part by two grants from the Ministry of Health, Labor and Welfare of Japan: the Research Project for Emerging and Reemerging Infectious Diseases (2001-2003): Molecular Analyses of Drug-Resistant Bacteria and Establishment of Rapid Identification Methods (grant no. H12-Shinkou-19), and Surveillance of Infectious Diseases Caused by Drug-Resistant Bacteria (grant no. H12-Shinkou-20).

 
* Corresponding author. Mailing address: Medical Microbiology Laboratory, Funabashi Medical Center, 1-21-1 Kanasugi, Funabashi, Chiba 273-8588, Japan. Phone: 81-47- 438-3321. Fax: 81-47-438-7323. E-mail: naganoyn{at}d3.dion.ne.jp.

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Journal of Clinical Microbiology, December 2003, p. 5530-5536, Vol. 41, No. 12
0095-1137/03/$08.00+0     DOI: 10.1128/JCM.41.12.5530-5536.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

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