Outbreak: the changing epidemiology of HAIs and CAIs

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LEARNING OBJECTIVES

Upon completion of this article, the
reader will be able to:

1. define and compare HAI and CAI;
2. discuss the three outbreak presentations and their significance;
3. review laboratory procedures to identify MRSA, A baumannii, and K pneumoniae;
4. review susceptibility-testing procedures and treatment; and
5. discuss changing patterns of epidemiology for each presentation.

T

he changing epidemiology of hospital-acquired (HA); nosocomial infection) and community-acquired (CA) infections has blurred the definition of these diseases. The present term, “healthcare-associated” infection (HAI) refers to disease appearing in any healthcare facility (hospital, nursing home, or outpatient clinic) >72 hours after admission. Predisposing risks for HAI include IV drug abuse, chronic or immune-resistant disease, advanced age (>65), previous hospitalization (or long-term-care facility [LTCF] ) residence), previous antibiotic therapy, and presence of an indwelling medical device or catheter.

“Community-associated” infection (CAI) describes a younger, healthier population in the community, who may experience toxic effects from invasive disease but who are without HAI risks. Exposure predominates in any community group — school children, athletic activities, military bases, and prisons — occurring 1,2

Since its appearance in the 1980s, distinguishing CA-MRSA (methicillin-resistant Stapholycoccus aureus) from HA-MRSA has led to the necessity for both phenotypic and genotypic characterization. Phenotypic identification is based on colonial features, biochemical identification, and antibiotic-susceptibility pattern. Genotypic patterns are determined technologically by the presence of a variety of other factors. In the case of MRSA, they include methicillin-resistance gene, SCCmec [SCCmec I-III for HA-MRSA, and SCCmec IV, V, VI (rare) for CA-MRSA], of clone groups (USA100, 200, or 500 for HA; USA300 or 400 for CA) and presence of virulence factors in CA-MRSA to explain toxicity.

Formerly, culture reports of patients infected 48 hours after admission to a healthcare facility presumed exposure to a nosocomial organism had occurred at the facility. Present findings make that assumption erroneous. Any of the “superbug” organisms can spread among susceptible patients and caregivers from exposure to colonized or infected persons in healthcare or community-type settings. Because of unique multidrug resistant (MDR) features, molecular characterization is needed for identification and treatment, and to provide surveillance data for infection control.¹

Gram negatives versus Gram positives

Escalating from penicillin-resistant S aureus in the 1940s to MDR and extensive-drug resistant tuberculosis (MDR-TB and XDR-TB) in the 1990s, resistance has followed the development of every antibiotic discovery. The beta lactams, cephalosporins, glycopeptides, fluoroquinolones, and carbapenems all have failed to eradicate infection with Gram-positive and Gram-negative superbugs (e.g., MRSA, Clostridium difficile, extended-spectrum beta lactamases [ESBLs] among the Enterobacteriaceae, and the non-fermentative Acinetobacter baumannii and Pseudomonas aeruginosa).

Having developed complex defenses against antibiotic therapy, MDR Gram-negative bacteria have reached formidable proportions in healthcare centers. An antibiotic approaching a bacterial cell encounters two cell walls, the second usually impenetrable. Another deterrent appears in the super-production of ESBL enzymes (e.g., Klebsiella pneumoniae, Escherichia coli, et al). ESBLs are derived from the TEM-1, SHV-1, or OXA enzymes and carried on plasmids with resistance genes for cephalosporins and aztreonam as well as the penicillins. The non-fermentative MDR bacilli and former environmental contaminant pests — Pseudomonas aeruginosa and A baumannii — have now reached significant proportions as causes of HAI and CAI with even greater potential for resistance. Like MRSA, these organisms usually require both phenotypic and genotypic work-up for complete characterization.

To illustrate the havoc evoked in both hospital and community by MDR organisms, three outbreak reports will be presented. Demonstrating the complexity of diagnosis, treatment, and prevention, they are CA-MRSA, A baumanni, and carbapenem-resistant K pneumoniae (KPC).¹

Outbreak #1: MRSA

A major threat to hospital and community is the notorious MRSA — primary cause of skin and soft-tissue infection with potential for invasive disease (e.g., blood, lung, cerebrospinal fluid [CSF]). A retrospective study (1997 to 2006) in a San Francisco LTCF examined the changing molecular epidemiology of MRSA genotypes. During the period from 1997 to 2003, the predominant isolate was the HA-MRSA clonal group USA100, an MDR strain (i.e., resistant to all beta-lactam drugs and resistant to >3 non-beta-lactam drugs [e.g., gentamicin, clindamycin, trimethorprim/sulfamethoxazole]). From 2004 to 2006, the CA-MRSA clone — USA300 — emerged. These strains were resistant to all beta lactams but had low resistance to non-beta-lactam antibiotics, with the exception of tetracycline. Fortunately, no resistance was reported to drugs of choice: vancomycin, linezolid, or daptomycin. Following the appearance of USA300 in the LTCF, the incidence of CA-MRSA isolates increased twofold, while HA-MRSA also increased, raising the overall incidence of infection from 38% to 73%.³

Laboratory identification of MRSA: Specimens from skin, soft-tissue, respiratory, wound, urinary-tract, and sterile sites (e.g., blood, lung, CSF) are collected, transported, and processed according to the protocol for Gram-positive organisms in the Clinical Manual of Microbiology.  

Culture media used primarily for growth includes sheep blood agar (SBA) and chocolate agar. Selective agars to inhibit Gram-negative organisms include phenylethyl alcohol agar (PEA), colistin-nalidixic acid agar,  and lipase-salt mannitol agar. Chromogenic agars (BD, Sparks, MD, and bioM’erieux, Hazelwood, MO) as well as oxacillin-salt agar are for the selective isolation of MRSA.^4,5

In Outbreak #1, 661 isolates of S aureus were screened with oxacillin-salt agar by inoculating Mueller-Hinton agar impregnated with 6 ug/mL of oxacillin and 4% NaCL. Similar to the disk-diffusion test, colonies are suspended in a broth (e.g., trypticase-soy or saline) adjusted to a 0.5 McFarland standard. The plate is swabbed in an area 10 mm to 15 mm in diameter, streaked on a quadrant of the agar surface, or spotted in the same area with a 1 uL loop.^4,5

Recommended only for S aureus to detect MRSA, the oxacillin-salt agar screen lacks sensitivity to detect borderline resistance (e.g., MIC <=4 ug/mL). Methods with greater sensitivity include rapid latex agglutination and the disk-diffusion cefoxitin test. (Note: The cefoxitin disk-diffustion test detects borderline strains that have the mecA gene; breakpoints are <=21 for resistant and >=22 for susceptible. In questionable cases, molecular testing can be performed by polymerase chain reaction [PCR]).^6,7 FDA-approved latex tests are the MRSA Screen Test (Denka-Seikin Co. Ltd., Tokyo, Japan) and the PBP 2 latex agglutination test (Oxoid Limited, Basingstoke, U.K.)

Antibiotic-susceptibility testing: Susceptibility testing follows the Clinical and Laboratory Standards Institute (CLSI) protocol, which recommends disk diffusion, broth, or agar dilution. Automated system testing, routinely used in most laboratories, is not valid for testing glycopeptides (e.g., vancomycin). Recommended are agar dilution tests (e.g., E Test, AB Biodisk, Solna, Sweden), capable of detecting vancomycin-intermediate or vancomycin-resistant S aureus.^6,7

Despite any susceptible in-vitro results, there are no beta-lactam drugs effective (in-vivo) to treat MRSA infection. CLSI mandates reporting methicillin-/oxacillin-resistant staphylococci resistant to all beta-lactam drugs — penicillins, cephalosporins, cephamycin, and  beta-lactam/beta-lactam inhibitors and carbapenems.^6,7

In Outbreak #1, diagnosis of MRSA was confirmed with PCR identification of the mecA gene; susceptibility testing was performed by microdilution with the MicroScan WalkAway 96 system (Dade Behring, Deerfield, IL). On isolates from 2005-2006, inducible clindamycin resistance was screened using the D-Zone Test, described and interpreted in CLSI, M7-A5 guidelines.^6,7

Epidemiology — molecular characterization: After DNA chromosomal digestion with genes SmaI, spa typing, and multilocus sequence typing (MLST), pulsed field gel electrophoresis (PFGE) genotyping was performed on the MRSA isolates to separate clone groups. PCR technology determined the presence of the Panton-Valentine leukocidin (PVL) toxin genes, lukF-PV and lukS-PV I, and the arginine catabolic mobile element — factors associated with CA-MRSA clone group USA300. The staphylococcal cassette chromosome mec type (SCCmec type IV) was identified by PCR; unidentifiable strains were subjected to MLST.³ (Note: Conflicting reports exist of PVL as a major virulence factor. The phagocytic role in skin and soft-tissue injury persists, while researchers consider other mechanisms involved in invasive disease.⁸)

Changing patterns: Epidemiologic tracking of MRSA noted two specific changes with the appearance of CA-MRSA: an emerging less-resistant antibiotic pattern and variation in specimen source. HA-MRSA clones are characterized by multiple drug resistance beyond the beta-lactams to include many non-beta-lactams, primarily in specimens isolated from urinary and respiratory tracts. CA-MRSA isolates are susceptible to most non-beta-lactam drugs and commonly isolated from skin and soft-tissue infection.

Another change noted in the San Francisco study was an unexpected increase in non-beta-lactam resistance in the CA-MRSA (USA300) clones. Statistically, a rising incidence occurred from 11% in 2002 to 64% in 2006. The increase led researchers to investigate the transmission rate in patients transferred from an LTCF to a hospital or community setting and back. Their findings determined that >30% of previously susceptible USA300 isolates became multidrug resistant during that period. Although drugs of choice continued to be susceptible (e.g., vancomycin, linezolid, dalbavancin, and daptomycin), resistance to gentamicin, clindamycin, trimethoprim-sulfamethoxazole, and tetracycline suggested selective resistance occurred in the LTCF, during residence in another hospital or facility or from a community-acquired contact.³

Managing infection-control policies in healthcare centers pales in comparison to instituting these programs in an LTCF. The logistics required in the isolation of positive patients, monitoring antibiotic-drug use, overseeing hand washing, and environmental cleaning in an LTCF is daunting. ³

Outbreak #2: Acinetobacter baumannii

Two decades ago, Acinetobacter spp. was well-recognized as a hospital contaminant. But the recent appearance of the MDR pathogen, A baumannii, has created a menacing environmental threat to both hospital and community. Tolerance of extreme temperatures and humidity as well as survival on dry surfaces for extended periods make these Gram-negative, non-fermentative opportunists ready contaminators of medical devices and equipment, (e.g., ventilator tubing, catheters, multidose medication vials, humidifiers).⁹

To study the epidemiologic features of nosocomial outbreaks in Los Angeles County, researchers acquired 20 clinical isolates of A baumannii, identified by the County Public Health Laboratory over an eight-year period (1996-2004). Correlation studies of phenotypic and genotypic features included analysis of antibiotic-susceptibility patterns, genomic fingerprints, and sequencing the quinolone resistance-determining region (QRDR) of the genome, performed to investigate the extent of fluoroquinolone resistance.¹⁰

Laboratory identification (Acinetobacter spp.): Specimen collection follows the protocol for Pseudomonas spp., described in the Clinical Manual of Microbiology. Respiratory specimens — sputum, lung fluid, or tissue — and urinary-tract and wound specimens are typical. After processing, aerobic culture plates (e.g., sheep blood agar [SBA], CA, MacConkey’s [MAC], and PEA) are inoculated with patient specimen. Anaerobic media (described in Outbreak #1) would be added when indicated (e.g., wounds, surgical specimens). Growth at 35^0C to 37^0C initially may be poor, requiring incubation of subculture and identifications tests to be carried out at <=30^0C.^4,11 Gram stains usually reveal Gram-negative bacilli or coccobacilli. Both staining and morphology are media dependent (e.g., in blood-culture media, the organism may deceive as Gram-positive cocci).

Strictly aerobic, non-motile, non-fermentative, oxidase negative, catalase positive (usually), and nitrate negative, A baumannii is commonly identified with commercial kits, as in Outbreak #2 (API 20 NE, bioM’erieux, Durham, NC.). With the addition of growth temperature testing at 44^0C, the kits replace more complex standard biochemical testing. To ensure accurate identification, molecular rRNA gene sequencing is considered state of the art and the clinical laboratory method of the future.¹¹

Antibiotic-susceptibility testing: Susceptibility testing for Acinetobacter spp., required for all clinically significant strains, presents problems of growth and interpretation. Standard methods of broth microdilution and disk diffusion may be discrepant for certain antibiotics (e.g., beta lactam and beta-lactam inhibitors). Although the Centers for Disease Control and Prevention (CDC) declines to indicate a preference for either method, CLSI has published breakpoints for both methods (tigecycline excepted).⁷

In Outbreak #2, the Los Angeles Public Health Laboratory chose 17 antibiotics and used the broth microdilution method with modifications, in accordance with CLSI. The antibiotics tested were amikacin, gentamicin, tobramycin, imipenem, meropenem, piperacillin, cefepime, cefotaxime, ceftazidime, ceftriaxone, ciprofloxacin, doxycycline, minocycline, gatifloxacin, levofloxacin, tetracycline, and tigecycline.^6,7,10

Antibiotic solutions were serially diluted with 5% dimethyl sulfoxide to microtiter plates (Costar 3795, Thermo Fisher Scientific, Tustin, CA), transferred to replicate plates, and covered in plastic to reduce evaporation. Plates were incubated at 35^0C for 18 to 24 hours and read visually with an inverted mirror to detect growth at the bottom of the wells. The lowest concentration without growth was defined as the minimal inhibitory concentration (MIC).¹⁰ Results were based on MIC ranges, designated: susceptible, intermediate, or resistant. Eighty percent of the isolates were susceptible only to tigecycline, doxycycline, and minocycline; 45% to 50% resistant to carbapenems (imipenem and meropenem); all isolates were resistant to ciprofloxacin (a fluoroquinolone). Comparison was made to five additional control organisms and American Type Culture Collection, or ATCC, reference strains.^6,7,10 Although study results showed tigecycline (MIC <=4 ug/mL) effective against 80% of the isolates, later reports demonstrated antibiotic resistance was increasing in vivo.¹⁰

Epidemiology — molecular characterization: Genomic DNA restriction digestion of the isolates was followed by the PFGE method described by Peleg and colleagues (Invitrogen, Carlsbad, CA) for analysis. A baumannii genes — gyrA and parC — were amplified and sequenced with PCR primer techniques (Applied Biosystems, Foster City, CA). Combined phenotypic and genetic features identified eight distinct epidemiologic lineages for the 20 isolates. All isolates were found to be multidrug resistant and highly related.¹⁰

In previous studies, 45% of both intensive-care unit (ICU) and non-ICU isolates were resistant to ciprofloxacin. Because 100% of the isolates in this study were resistant, a search for mechanisms of resistance was undertaken. Two were hypothesized: a change in drug-target protein structure and presence of efflux pumps, capable of removing fluoroquinolones from the cell — a phenomenon known to other Gram-negative bacteria. Sequencing the 20 isolates revealed point mutations on the gyrA or parC genes had resulted in amino-acid substitutions. This discovery determined that changes in the protein target, leaving A baumanni resistant to fluoroquinolones, indeed, had occurred. Susceptibility testing performed in the presence and absence of efflux pump inhibitors PAN or NMP (added to the microplate wells) confirmed these were not major contributors to the ciprofloxacin resistance. Evidence of point mutations in the QRDRs (gyrA and parC genes) supported the theory of selective antibiotic pressure in a hospital environment.¹⁰

Changing patterns: Increased rates of catheter-associated bloodstream infection, ventilator-associated pneumonia (VAP), or meningitis, and urinary-tract and wound infections comprise 2% to 10% of Gram-negative infections in the ICUs of the U.S. and Europe. Although infection caused by A baumannii in the community is limited, mortality rates when community-acquired pneumonia is present have been reported as high as 40% to 64%.⁹

Reports of military casualties from Iraq and Afghanistan indicate A baumannii is the most common isolate from “war wounds,” with strains resistant to the major classes of antibiotics: beta lactams and inhibitors, aminoglycosides, fluoroquinolones, cholamphenicol, tetracycline, and rifampin.⁹

Epidemiologic studies have suggested that fecal surveillance in the hospital environment may reduce outbreaks, particularly in the high risk area of ICU, where VAP is common. To reduce contamination from hospital rooms, bedding, and other furnishings and deter patient colonization, however, requires regular stringent cleaning with sodium hypochlorite solution (1000 pp/m).⁹

OUTBREAK # 3 Klebsiella pneumoniae, carbapenem-resistant (KPC)

As a class, the carbapenem antibiotics (e.g., imipenem, meropenem) have been effective in treating disease caused by ESBL-producing Gram-negative bacteria. ESBL enzymes have increased potency against cephalosporin antibiotics, resulting from critical amino-acid substitutions during bacterial mutation derived from common beta-lactamase genes (e.g., TEM-1, SHV-1, and CTX-M, in K pneumoniae, Klebsiella oxytoca, Escherichia coli, and others) The unique capability of the first two is hydrolysis of penicillins and some cephalosporins; while CTX-M is active against cefotaxime and ceftriaxone (second-generation cephalosporins), it is not active agaomst ceftazidime (third-generation cephalosporin).^1,12

An isolate of the KPC-producing bacteria was first reported in North Carolina (1996), then serious outbreaks were found in New York City and New Jersey (2001), and finally it spread globally. In January 2007, an outbreak occurred in a Virginia 310-bed community hospital, auspiciously noted by a laboratory manager, who observed an increase in ESBL isolates with elevated imipenem MICs. His report prompted an investigation by the Duke Infection Control Outreach Network, which found 58 patients with probable KPC had been admitted as transfers from nursing homes and LTCFs. The case definition of a probable KPC isolate stated that ESBL bacteria have an MIC >=1 ug/mL to imipenem by broth dilution, confirmed by the Modified Hodge Test (a disk-diffusion procedure, using meropenem or ertapenen discs on Mueller-Hinton agar to detect carbapenemase production).^13,14

Twenty-nine isolates of probable KPC were sent to Israel for molecular typing by PFGE. Results determined only two genotypes (KPC2 and KPC3) were present in the outbreak, and none had identical fingerprinting despite their high clonal-relatedness, defined by a 76% homology between type V and the others. In other words, patients from the same nursing home had the same genotype. Similar KPC types supported other findings to indicate that transmission occurred from exposure in both hospital and community environment.

Significant information derived from surveillance in Outbreak #3 revealed that >50% of the patients had community-acquired infection according to the standard definition (i.e., 13

Laboratory identification of KPC: Specimen collection and preparation for detecting K pneumoniae follows the protocol for Enterobacteriaceae outlined in the Clinical Manual of Microbiology. Specimens from respiratory, urinary-tract, wound, and sterile sites are submitted to the laboratory and processed for aerobic culture on SBA, PEA, and MAC. When indicated, anaerobic media are added (e.g., brucella blood agar [BBA], anaerobic PEA, and BBA with gentamicin, or BG).^4,15

Recently developed for ESBL isolation are two selective chromogenic agars, Brilliance ESBL agar, (OX; Oxoid, Basingstoke, U.K.), and ChromID ESBL agar (BM; bioMerieux, Marcy l’Etoile, France). When compared to MAC supplemented with cefotaxime and/or ceftazidime, the selective agars proved to have higher sensitivity and specificity for the presumptive identification of ESBLs.^15,16

K pneumoniae is a facultative, fermentative Gram-negative rod, identifiable with standard biochemicals, by manual (kit) identification system, (e.g., API  50 CH, API ZYM carbohydrate and enzymatic panels, bioMerieux, Hazelwood, MO) or by automated system (e.g., VITEK, bioMerieux, Marcy l’Etoille, France; Dade MicroScan, Dade Behring, Deerfield, IL, et al.)

The slow growth and temperature dependence common to some Klebsiella spp., however, may result in detection error with automated system identification. Standard biochemical tests also have demonstrated a lack of reliability without attention to low temperature growth and carbon assimilation. Growth at 10^0C, histamine utilization and lack of gas from lactose fermentation at 44.5^0C differentiates Raoultella spp. from K pneumoniae and K oxytoca. A newly discovered species, K varicola, is separated from K pneumoniae only by lack of adonitol fermentation.^13,15

Because of the unique antibiotic-resistance pattern involving cephalosporin activity, the correct identification of KPC is imperative. Adding to the treatment dilemma posed by ESBL production is decreased susceptibility to the fluoroquinolone antibiotics exhibited by K pneumoniae and K oxytoca (7% and >3%, respectively).¹⁵

Antibiotic-susceptibility testing: Susceptibility testing follows the CLSI protocol for fermentative, Gram-negative bacilli (i.e., disk diffusion, broth or agar dilution, and automated system testing). KPC-producing organisms, presenting detection problems in automated systems, may not “flag” as ESBL if susceptibility to second-generation cephalosporin agents occurs. In these cases, lower testing dilutions will be required to detect carbapenem resistance. CLSI provides breakpoints for screening aztreonam, cefotaxime, cefpodoxime, ceftazidime, and ceftriaxone for all ESBL-producing organisms when disk diffusion or broth microdilution is performed. CLSI recommends clinical laboratory reports of ESBLs should state: “resistant to all extended-spectrum penicillins, cephalosporins, and monobactams” to ensure successful in vivo treatment. When reliable results are not available, reference, and public-health laboratories will provide PCR, sequencing technology, and PFGE fingerprinting as in Outbreak #3.^6,7

Poor detection of KPC-producing bacteria by automated systems occurs because of low sensitivity and specificity for the carbapenemase enzyme when antibiotics imipenem or meropenem are tested. The most reliable confirmation of carbapenem resistance is provided by testing for the bla_KPC gene with PCR technology. In the KPC isolates from Outbreak #3, the Modified Hodge Test (M100-S19 CLSI January 2009) was used to confirm carbapenem resistance.^7,14 (Note: The 2010 CLSI committee revised and published new carbapenem breakpoints in June 2010, negating the need for the Modified Hodge Test.⁷)

The high mortality rate in ICU patients (22% to 59%) necessitates effective treatment with the less-common polymixin antibiotics. Though noted for lower nephrotoxicity than previously believed, reports of increasing resistance occur. As is generally true, in Outbreak #3, tigecycline was reported to have exquisite susceptibility in cases of KPC. Although no CLSI breakpoints exist for tigecycline, any KPC resistance by standard microdilution methods is presumed minimal.¹³

Epidemiology: Reported by 27 states in the U.S., carbapenemase enzymes (e.g., beta lactamases) are spreading rapidly throughout the world. Prior to several outbreaks in the metropolitan areas of New York and New Jersey in 2001, however, isolation and reporting were rare. Usually detected in Klebsiella spp. and E coli, KPC bacteria are reported in many Enterobacteriaceae and in MDR Pseudomonas aeruginosa.

The epidemiology of KPC genes continues to be investigated. While carried on plasmids capable of jumping-transfer among other bacterial strains and species, they hydrolyze and render ineffective the activity of penicillins, cephalosporins, and carbapenems.¹³

Screening: As selective pressure from broad-spectrum antibiotics has led to colonization of the digestive and respiratory tracts of susceptible patients in nursing homes and hospitals, transmission of KPC genes has become a major concern.¹³ Previous strategies to control ICU outbreaks included culture-screening methods for new patient admissions by direct plating of rectal swab or stool samples to MAC agar with imipenem and ertapenem discs and 24-hour incubation. The PCR-based method to detect colonization of KPC-producing Klebsiella species, however, has higher sensitivity than culture — 96% versus 78% — and shorter turnaround time — 30 hours versus 60 to 75 hours.¹⁷

Changing patterns: CDC and the Healthcare Infection Control Practice Advisory Committee (HICPAC) 2009 guidelines for treatment of KPC advocated aggressive action to deter its spread. No new antibiotics are available with a mechanism of action to deter production of the unique jumping-transfer of KPC carbapenemases, which create low-level resistance in the major antibiotics chosen to treat those infections. The confirmed increase in KPC resistance (east to west) and risk of necrotizing infection mandated an immediate prevention protocol: 1) determine source of the carbapenem resistance by review of microbiology records (six to 12 months); 2) perform point-prevalence culture survey in high-risk areas to look for other cases; and 3) perform active surveillance cultures of patient contacts.¹⁸

Conclusion

A number of antibiotic-resistant organisms, the cause of hospital-acquired infection, now appear to threaten the community. Excessive use of antibiotics (selective antibiotic pressure) has created an alarming increase in MDR diseases. With added potential for severe toxicity, CA-MRSA infection exemplifies the high mortality rate associated with invasive disease. A baumannii, a rare but morbid cause of HAI or CAI pneumonia, ultimately may be untreatable. Most significant is that rapid transmission of difficult-to-detect KPC genes in hospital and community settings has become a worldwide threat.

As demonstrated in the three cases presented, treatment options are limited, antibiotic choices minimal, and the need for fast diagnosis by the clinical laboratory critical. Molecular techniques such as 16S rRNA sequencing for difficult-to-identify organisms (e.g., Acinetobacter spp.) and real-time PCR testing of resistant genes (e.g., ESBLs, Amp C beta lactamases, carbapenemases) can provide earlier and better-targeted empiric therapy. Adopting real-time surveillance of prevalent beta-lactamase genes in a particular healthcare center also shows added promise for success in the war against MDR bacteria.¹⁹

Prevention and control of MDR organisms: Strategies to manage MRSA, A baumannii, and KPC require aggressive management: stringent healthcare hygiene, active surveillance, antibiotic stewardship, and application of epidemiologic methods for the particular organism. Members of a stewardship team can maximize efficacy by increasing dosage or adjusting infusion time to minimize antibiotic exposure. Combining the expertise of the microbiology laboratory and information technology can provide digital surveillance to prevent poor antibiotic choices. Finally, application of molecular methodology, not simply as a research tool but in the clinical laboratory, will assure accurate identification and faster diagnosis.²⁰

Cynthia B. Schofield, MPH, MT(CAMT), is a microbiology technical supervisor (retired) from the VA San Diego Healthcare Systems in San Diego, CA.

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