JOURNAL OF CLINICAL MICROBIOLOGY,
Copyright © 1999, American Society for Microbiology. All Rights Reserved.
Aug. 1999, p. 2479–2482Vol. 37, No. 8
PCR and Blood Culture for Detection of Escherichia coli
Bacteremia in Rats
ALEXANDRA HEININGER,1* MARLIES BINDER,2SIBYLLE SCHMIDT,2KLAUS UNERTL,1
KONRAD BOTZENHART,2AND GERD DO ¨RING2
Klinik fu ¨r Ana ¨sthesiologie1and Department of General and Environmental Hygiene,2Hygiene Institute,
University of Tu ¨bingen, Tu ¨bingen, Germany
Received 5 January 1999/Returned for modification 19 March 1999/Accepted 20 April 1999
Critically ill patients often develop symptoms of sepsis and therefore require microbiological tests for
bacteremia that use conventional blood culture (BC) techniques. However, since these patients frequently
receive early empirical antibiotic therapy before diagnostic procedures are completed, examination by BC can
return false-negative results. We therefore hypothesized that PCR could improve the rate of detection of
microbial pathogens over that of BC. To test this hypothesis, male Wistar rats were challenged intravenously
with 106CFU of Escherichia coli. Blood was then taken at several time points for detection of E. coli by BC and
by PCR with E. coli-specific primers derived from the uidA gene, encoding ?-glucuronidase. In further
experiments, cefotaxime (100 or 50 mg/kg of body weight) was administered intravenously to rats 10 min after
E. coli challenge. Without this chemotherapy, the E. coli detection rate decreased at 15 min and at 210 min after
challenge from 100% to 62% of the animals with PCR and from 100% to 54% of the animals with BC (P, >0.05).
Chemotherapy decreased the E. coli detection rate at 25 min and at 55 min after challenge from 100% to 50%
with PCR and from 100% to 0% with BC (P, <0.05). Thus, at clinically relevant serum antibiotic levels, PCR
affords a significantly higher detection rate than BC in this rat model. The results suggest that PCR could be
a useful adjunct tool supplementing conventional BC techniques in diagnosing bacteremia.
Episodes of sepsis are common and frequently life-threat-
ening (12) complications, occurring in 44.8% of all critically ill
patients (38). By definition (4), sepsis is caused by microbial
infection. However, bacteremia, i.e., the presence of microor-
ganisms in the bloodstream, is diagnosed in only approximately
4 to 12% of all blood cultures (BC) (26, 37). Escherichia coli is
one of the most common microorganisms isolated during hos-
pital-acquired bacteremia in intensive care patients (2, 21, 38).
Several factors may contribute to the low bacteremia detection
rate. First, some species of bacteria are difficult to culture (9),
and some, such as Mycobacterium spp. (5, 11, 15) or Bartonella
spp. (6), grow very slowly. Second, bacteremia is often tran-
sient (29), meaning that the number of viable microorganisms
circulating in the blood decreases rapidly after the onset of
bacteremia due to phagocytic and other host defense mecha-
nisms. Finally, up to 65% of intensive care patients showing
symptoms of sepsis are already under antimicrobial treatment,
thus compromising the results of microbiological culture tech-
niques (8, 25, 31, 37).
Since the emergence of bacteremia is associated with high
mortality rates and financial costs as well as a longer hospital
stay (7, 27, 30), independent of the severity of the underlying
disease (3, 33), improved techniques for diagnosing bacteremia
are needed. We hypothesized that amplification of bacterial
DNA by PCR could overcome some of the problems associ-
ated with conventional BC techniques. The potential of PCR
technology to recover intracellular or nongrowing microorgan-
isms may be useful for diagnosing bacteremia. Although PCR
has already been successfully applied in the detection of fas-
tidious microorganisms (22, 43) and, in particular, for detect-
ing bacteremia (23, 28, 44), a direct comparison of BC tech-
niques with PCR with respect to detection levels and time
course during bacteremia has yet to be carried out. Therefore,
we assessed these parameters for experimental bacteremia in-
duced by E. coli in rats.
MATERIALS AND METHODS
Rat model of bacteremia. Male Wistar rats (350 to 500 g) were anesthesized by
an initial intramuscular injection of ketamine at 80 mg/kg of body weight (bw)
and xylazine at 4 mg/kg of bw, separately. Following this injection, both jugular
veins were cannulated with a 26-gauge cannula (Abbott, Wiesbaden, Germany)
under aseptic conditions. One cannula was used exclusively for the collection of
blood samples. Bacteremia was induced via the other cannula by injection of 100
?l of a suspension of E. coli (ATCC 11229; 107CFU/ml in 0.9% saline). E. coli
was chosen to represent nonfastidious pathogens for comparing the sensitivities
of BC and PCR because it is the most frequent gram-negative pathogen causing
bacteremia. We collected 2.2-ml blood samples from 19 animals (group I) 5 min
before bacterial challenge and at 5, 15, 150, 180, and 210 min afterward. One
blood sample from each of 16 animals surviving 210 min was analyzed at each
time point, with the exception of time points 15, 150, and 210 min after bacterial
challenge, when analysis for E. coli was limited to 10, 15, and 13 rats, respectively,
due to insufficient blood sample volumes. One milliliter of whole blood was
immediately inoculated into the BC medium, and 1 ml was stored in EDTA tubes
(Sarstedt, Nu ¨rmbrecht, Germany) for PCR analysis; 100 ?l of each sample was
directly plated in duplicate on sheep blood agar plates for the quantitation of
From an additional 20 animals, 2.0-ml blood samples were collected before
bacterial challenge and at 5, 25, 40, and 55 min afterward for detecting E. coli by
BC and PCR. Ten animals (group II) received 100 mg of cefotaxime/kg of bw 10
min after bacterial challenge via the cannula used to inject E. coli. The other 10
animals (group III) received 50 mg of cefotaxime/kg of bw. At each time point,
one blood sample was analyzed from each of 10 animals. From three animals in
each group, an additional 0.5 ml of blood was taken and stored in an EDTA tube
(Sarstedt) to check serum cefotaxime concentrations. The experiments were
approved by the Animal Ethics Committee of the University of Tu ¨bingen, Tu ¨-
In order to detect E. coli by BC, 1-ml blood samples were inoculated into
BACTEC Peds Plus F medium (Becton Dickinson, Sparks, Md.), which contains
resin for neutralizing antimicrobial agents (13, 31, 45) and is designed for an
inoculum of 1 to 3 ml. The vials were incubated for 7 days in a shaker incubator
at 37°C. After 48 h, the vials were punctured under sterile conditions, and 100 ?l
was subcultured on sheep blood (5%) agar in duplicate for 24 h (5% CO2). If
bacterial growth was negative, the vials were incubated for another 5 days,
followed by subculturing as described above.
* Corresponding author. Mailing address: Klinik fu ¨r Ana ¨sthesiolo-
gie, University of Tu ¨bingen, Hoppe-Seyler-Str. 3, D-72076 Tu ¨bingen,
Germany. Phone: 0049 7071 2986622. Fax: 0049 7071 295533. E-mail:
PCR. A 1-ml blood sample was divided into three aliquots of 300 ?l each,
which were transferred into sterile cups. Erythrocytes were lysed, and DNA was
extracted according to the manufacturer’s protocol with a Puregene DNA iso-
lation kit (Biozym Diagnostik GmbH, Hessisch Oldendorf, Germany) for whole
blood. Two pairs of oligonucleotide primers (one nested in the other) were
derived from the uidA gene of E. coli, encoding ?-glucuronidase specific for E.
coli and Shigella spp. Oligonucleotides P1 (5?-ATC ACC GTG GTG ACG CAT
GTC GC-3?) and P2 (5?-CAC CAC GAT GCC ATG TTC ATC TGC-3?) were
used in the first round of PCR to amplify a 486-bp fragment (19). Oligonucle-
otides P3 (5?-TAT GAA CTG TGC GTC ACA GCC-3?) and P4 (5?-CAT CAG
CAC GTT ATC GAA TCC-3?) were used in the nested PCR for the amplified
products from the primary PCR to amplify a 186-bp fragment. For DNA ampli-
fication, 20.4 ?l of reaction mixture was added to 29.6 ?l of purified DNA. The
master mixture for the first round of PCR included the following: 10 mM
Tris-HCl, 50 mM KCl, 4.5 mM MgCl2, 600 ?M each deoxynucleoside triphos-
phate (dNTP) (Amersham Pharmacia Biotech, Freiburg, Germany), 0.6 ?M
each primer, and 1 U of Taq DNA polymerase (Amersham Pharmacia Biotech).
Each reaction mixture was overlaid with 1 or 2 drops of mineral oil. PCR
amplifications were performed with a Hybaid Omni Gene Temperature Cycler
(MWG Biotech, Ebersberg, Germany).
The primary PCR amplification consisted of an initial denaturation step at
95°C for 5 min; 35 cycles at 95°C for 30 s, 50°C for 1 min, and 72°C for 1 min; and
a final elongation phase at 72°C for 5 min. After the first reaction, 1 ?l of the
amplified product was added to 49 ?l of the second master mixture. The master
mixture for the nested PCR contained the following: 10 mM Tris-HCl, 50 mM
KCl, 4 mM MgCl2, 400 ?M each dNTP (Amersham Pharmacia Biotech), 0.4 ?M
each primer, 10 ?l of Q-solution (Qiagen, Hilden, Germany), and 1 U of Taq
DNA polymerase (Amersham Pharmacia Biotech). The nested PCR amplifica-
tion was performed with an initial denaturation step at 95°C for 5 min; 35 cycles
at 95°C for 30 s, 50°C for 1 min, and 72°C for 1 min; and a final elongation phase
at 72°C for 5 min.
Each time a PCR was performed, we also ran a positive control (bacterial
DNA), a negative control (rat DNA), and a reagent control (all PCR reagents
without DNA) to evaluate the success of amplification, the specificity of the
reaction, and the purity of the reagents, respectively. The product of the nested
PCR was electrophoresed with 2 ?l of gel loading buffer (0.25 g of bromphenol
blue and 40 g of saccharose dissolved in 100 ml of distilled water) through a 2%
agarose gel at 110 V for 40 min in Tris-acetate–EDTA buffer). Molecular size
markers (Promega, Madison, Wis.) were run concurrently. The gel, stained with
ethidium bromide (0.5 ?g/ml), was examined under UV light for the presence of
a 186-bp band and photographed for documentation.
In order to assess the sensitivity of a specific PCR, 1 ml of rat blood was spiked
with E. coli (106to 100CFU) and immediately processed as described above.
Verification of correct fragment production by the PCR was performed by
Southern hybridization. PCR fragments were Southern blotted onto positively
charged nylon membranes (Boehringer GmbH, Mannheim, Germany) by use of
a Turbo Blotter (Schleicher & Schuell, Inc., Keene, N.H.) as prescribed by the
manufacturer for alkaline transfer. The DNA was cross-linked to the damp
membranes by use of a GS Gene Linker UV chamber (BioRad Laboratories,
Hercules, Calif.) according to the instructions of the provider. The gene probe
used for hybridization was a digoxigenin-UTP-labeled (Boehringer) 186-bp frag-
ment produced by PCR, resolved on a 2% agarose gel, and purified with a PCR
product purification kit (Qiagen). The standard nylon membrane prehybridiza-
tion and hybridization protocol was performed under stringent conditions at
65°C. Probed membranes were exposed to Hyperfilm ECL (Amersham Life
Science, Little Chalfont, England).
Analysis of cefotaxime levels in serum. Cefotaxime concentrations were mea-
sured by high-performance liquid chromatography with a UV detection system
by the method reported by Dell et al. (10).
Statistical analysis. In order to detect any relevant difference between the
PCR and the BC methods with respect to bacteremia detection as a function of
time, we chose confidence intervals of 95% (P, ?0.05). The minimum number of
experiments needed to establish a difference of at least 30% between the two
techniques for group I was calculated to be 20. A preliminary data analysis was
scheduled after the completion of 12 experiments. For groups II and III, the
confidence intervals were also 95%. The minimum number of experiments
needed to reveal a difference of at least 80% between the two methods for groups
II and III was 10 (1).
For the detection of E. coli DNA in rat blood samples, PCR
was performed with primers for the uidA gene of E. coli (19).
PCR products from rat blood samples were verified to be
specific for the target gene by Southern hybridization with a
dUTP-labeled probe for uidA. No differences in the sizes of the
PCR fragments from rat blood samples on agarose gels and the
hybridization product on the Southern blot were observed
(data not shown).
The sensitivity of PCR and BC for detecting E. coli in whole
rat blood was 1 CFU/ml (data not shown). Before bacterial
challenge, BC, the direct plating method, and PCR all returned
negative results, indicating the absence of bacterial contami-
nation during the animal experiment and during the PCR
procedure (Fig. 1A). After bacterial challenge, the rates of
detection of E. coli by the three methods decreased in a time-
dependent manner (Fig. 1A). Even after 210 min, the E. coli
PCR was positive for 61% of the 13 blood samples, whereas
BC with the BACTEC system and the direct plating method
were positive for 53 and 8% of the samples, respectively. No
significant difference was observed between PCR and BC for
the detection of E. coli at any of the examination times, but the
rate of detection by the direct plating method was noticeably
lower than that by either PCR or BC. Quantitatively, the direct
plating method showed mean numbers of bacteria per milliliter
of 27, 15, 0.0, 0.6, and 0.4 at 5, 15, 150, 180, and 210 min after
E. coli challenge.
FIG. 1. Time course of E. coli detection in experimental bacteremia in rats by
PCR, BC, and the direct plating method. (A) Sixteen rats were challenged
intravenously with 106CFU of E. coli at time zero (arrow a) without further
intervention. (B and C) Another 10 rats each received 100 mg (B) (arrow b) or
50 mg (C) (arrow c) of cefotaxime/kg of bw 10 min after bacterial challenge. For
E. coli detection, blood was collected 5 min before challenge and at the indicated
times after challenge. E. coli was detected by PCR (A, B, and C) (filled bars), BC
(A, B, and C) (hatched bars), and the direct plating method (A) (open bars).
Bars represent the percentages of E. coli-positive blood samples. ?, P ? 0.05.
2480HEININGER ET AL. J. CLIN. MICROBIOL.
As expected, the sensitivity of BC decreased significantly
during experimental E. coli bacteremia in rats receiving clini-
cally relevant doses of cefotaxime (Fig. 1B and C). In antibiotic
treatment groups II and III, peak serum cefotaxime levels
ranged between 121 and 137 mg/liter and between 94 and 86
mg/liter 15 min after injection, respectively (data not shown).
When blood was taken 15 min after cefotaxime was adminis-
tered intravenously, i.e., 25 min after bacterial challenge, only
10% of the samples were found positive by BC, whereas 100%
of the samples were found positive by PCR (Fig. 1B and C).
Data analysis established an 80% higher detection rate for the
PCR method 25 min after bacterial challenge with both cefo-
taxime doses. At subsequent sampling times, the advantage of
the PCR technique was also evident but did not reach the 80%
BC is considered the “gold standard” for detecting micro-
organisms in the blood (29, 37). Nevertheless, many clinicians
and microbiologists are concerned that BC for critically ill
patients receiving antimicrobial chemotherapy could indicate
lower rates of bacteremia than are actually present (8, 34).
Antibiotic-binding BC devices, such as the BACTEC Plus F
system, were developed in response to these concerns (13, 31,
41). Still, the rate of detection of bacteria in blood specimens
has not changed substantially. The results of the present study
strongly suggest that the clinicians’ suspicions have a scientific
basis. In an experimental rat model for E. coli bacteremia, a
significantly higher rate of detection of E. coli was achieved
with PCR technology than with BC when the animals were
treated with clinically relevant levels of cefotaxime.
Interestingly, this difference in the detection rate between
PCR and BC was not due to a higher sensitivity of our PCR
than of BC. Regardless of whether PCR or BC is used, the
ability to detect bacteremia depends on the presence of at least
one microorganism in the blood being sampled. When tested
in vitro, both methods detected as little as 1 CFU of E. coli per
ml of blood, a volume which is normally used for diagnosing
bacteremia, particularly in neonatal intensive care units (26).
Furthermore, no significant difference in E. coli detection rates
was observed between PCR and BC at any time point during
experimental bacteremia in animals without antibiotic treat-
ment. The sensitivity of our PCR was comparable to that pre-
viously reported for PCR used to detect bacteremia (17, 23, 28,
32, 35, 44). To our knowledge, however, minimal detection
levels of BC systems in vitro have yet to be reported. Since the
number of pathogens is less than 1 microorganism per ml of
blood in 62% of all adult patients with E. coli bacteremia (37)
and can be less than 0.04 organism per ml of blood (18), large
blood volumes (20 to 30 ml) have to be used in order to avoid
false-negative results. Thus, even though the PCR method was
designed for smaller volumes from the very outset, it has to be
adapted accordingly. This problem might be solved in the near
future, because the rapidly increasing clinical use of PCR tech-
nology to detect pathogens which could not be isolated by
conventional methods (5, 15, 24) has already prompted the
development of a variety of different DNA isolation kits.
Our finding that BC is much less efficient than PCR in
detecting bacteremia during antimicrobial treatment is most
probably due to the killing of E. coli by cefotaxime; killed
bacteria are not detectable by BC, while PCR detects bacterial
DNA independently of viability.
Similarly, in a rabbit model of endocarditis, recovery rates
for E. coli and other bacteria were reduced during antimicro-
bial treatment (25). In addition, the higher sensitivity of PCR
technology than of conventional BC techniques is due to the
fact that intracellular or phagocytosed microorganisms are also
detectable by PCR.
In order to quantify bacterial numbers in E. coli-challenged
rat blood, the direct plating method was used. Direct plating
corroborated the results obtained by PCR and BC with respect
to the rapid clearance of E. coli from the bloodstream. The
lower sensitivity of the direct plating method compared with
PCR or BC may be due to its smaller sample volume and a lack
of dilution of bacterial growth-inhibiting factors in the blood,
which are known to influence bacterial recovery rates in BC
(29, 37). Thus, since direct plating is clearly less sensitive than
BC in detecting E. coli in blood samples, BC is a better stan-
dard than direct plating for the evaluation of PCR technology
From the clinical point of view, it is vital to develop im-
proved techniques for diagnosing bacteremia, because the
emergence of bacteremia is of substantial prognostic and ther-
apeutic importance (2, 16, 39). The occurrence of secondary
bacteremia (14) is a signal that the host’s defenses have failed
to contain an infection at its primary site or that the physician
has failed to control the infectious process (40, 42). Thus, the
detection of bacteria in the patient’s blood, regardless of
whether they are still viable or have been killed by antibiotics,
implies that the treatment regimen might be insufficient and
has to be augmented.
Extracellular bacterial DNA has been considered to be im-
munologically inert in mammals. However, it was shown re-
cently that bacterial DNA has substantial immunostimulatory
properties comparable to those of endotoxin (36) and that its
presence can cause sepsis-like symptoms in mice (36). Thus,
methods for detecting DNA might improve our understanding
of the frequently life-threatening systemic inflammatory re-
sponse syndrome (4) in intensive care patients. In summary,
our results suggest that PCR could prove to be a useful adjunct
tool supplementing conventional BC techniques in diagnosing
We are indebted to W. Hoffmann and T. Gottwald of the University
of Tu ¨bingen for technical assistance concerning the rat model; K.-J.
Plaueln of Hoechst-Marrion-Roussel, Frankfurt, Germany, for the
kind gift of cefotaxime; M. Ulrich, C. Goerke, P. Kru ¨ger, and C. Wolz
for discussions concerning the establishment of PCR technology; C.
Meisner for assistance in the statistical evaluation of our data; D.
Isaacman for reading the manuscript; and D. Blaurock for language
corrections in the manuscript.
1. Altman, D. G. 1991. Comparing groups—categorical data, p. 229–276. In
D. G. Altmann (ed.), Practical statistics for medical research. Chapman &
Hall, Ltd., London, England.
2. Bachur, R., and G. L. Caputo. 1995. Bacteremia and meningitis among
infants with urinary tract infections. Pediatr. Emerg. Care 11:280–284.
3. Bates, D. W., K. E. Pruess, and T. H. Lee. 1995. How bad are bacteremia and
sepsis? Arch. Intern. Med. 155:593–598.
4. Bone, R. C., R. A. Balk, F. B. Cerra, R. P. Dellinger, A. M. Fein, W. A. Knaus,
R. M. H. Schein, and W. J. Sibbald. 1992. Definitions for sepsis and organ
failure and guidelines for the use of innovative therapies in sepsis. Chest
5. Bottger, E. C., A. Teske, P. Kirschner, S. Bost, H. R. Chang, V. Beer, and B.
Hirschel. 1992. Disseminated Mycobacterium genevase infection in patients
with AIDS. Lancet 340:76–80.
6. Brenner, S. A., J. A. Rooney, P. Manzewitsch, and R. L. Regnery. 1997.
Isolation of Bartonella (Rochalimaea) henselae: effects of methods of blood
collection and handling. J. Clin. Microbiol. 35:544–547.
7. Crowe, M., P. Ispahani, H. Humphreys, T. Kelley, and R. Winter. 1998.
Bacteraemia in the adult intensive care unit of a teaching hospital in Not-
tingham, UK, 1985–1996. Eur. J. Clin. Microbiol. Infect. Dis. 17:377–384.
8. Darby, J. M., P. Linden, W. Pasculle, and M. Saul. 1997. Utilization and
VOL. 37, 1999 E. COLI DETECTION BY PCR OR BLOOD CULTURE2481
diagnostic yield of blood cultures in a surgical intensive care unit. Crit. Care
9. Davies, S., and R. Eggington. 1991. Recovery of Mycoplasma hominis from
blood culture media. Med. Lab. Sci. 48:110–113.
10. Dell, D., J. Chamberlain, and F. Coppin. 1981. Determination of cefotaxime
and desacetylcefotaxime in plasma and urine by high-performance liquid
chromatography. J. Chromatogr. 226:431–440.
11. Esteban, J., A. Molleja, R. Fernandez-Roblas, and F. Soriano. 1998. Number
of days required for recovery of mycobacteria from blood and other samples.
J. Clin. Microbiol. 36:1456–1457.
12. Fagon, J. Y., A. Novara, F. Stephan, et al. 1994. Mortality attributable to
nosocomial infections in the ICU. Infect. Control Hosp. Epidemiol. 15:428–
13. Fuller, D. D., and T. E. Davis. 1997. Comparison of Bactec plus Aerobic/F,
Anaerobic/F, Peds Plus/F and Lytic/F media with and without fastidious
organism supplement to conventional methods for culture of sterile body
fluids. Diagn. Microbiol. Infect. Dis. 29:219–225.
14. Garner, J. S., W. R. Jarvis, T. G. Emori, T. C. Horan, and J. M. Hughes.
1988. CDC definitions for nosocomial infections. Am. J. Infect. Control
15. Haas, W. H., P. Kirschner, S. Ziesing, H. J. Bremer, and E. C. Bottger. 1993.
Cervical lymphadenitis in a child caused by a previously unknown mycobac-
terium. J. Infect. Dis. 167:237–240.
16. Isaacman, D. J. 1998. Strategies for improving the detection of bacteremia in
children. Infect. Dis. Clin. Pract. 7:28–31.
17. Isaacman, D. J., Y. Zhang, J. Rydquist-White, R. M. Wadowsky, J. C. Post,
and G. D. Ehrlich. 1995. Identification of a patient with Streptococcus pneu-
moniae bacteremia and meningitis by the polymerase chain reaction (PCR).
Mol. Cell. Probes 9:157–160.
18. Jonsson, B., A. Nyberg, and C. Henning. 1993. Theoretical aspects of detec-
tion of bacteraemia as a function of the volume of blood cultured. APMIS
19. Juck, D., J. Ingram, M. Prevost, J. Coallier, and C. Greer. 1996. Nested PCR
protocol for the rapid detection of Escherichia coli in potable water. Can. J.
20. Kane, T. D., S. R. Johnson, J. W. Alexander, G. F. Babcock, and C. K. Ogle.
1996. Detection of intestinal bacterial translocation using PCR. J. Surg. Res.
21. Kiehn, T. E., B. Wong, F. F. Edwards, and D. Armstrong. 1983. Comparative
recovery of bacteria and yeasts from lysis-centrifugation and a conventional
blood culture system. J. Clin. Microbiol. 18:300–304.
22. Kox, L. F. F., D. Rhienthong, A. M. Miranda, N. Udomsantisuk, K. Ellis, J.
van Leeuwen, S. van Heusden, S. Kuijper, and A. H. J. Kolk. 1994. A more
reliable PCR for detection of Mycobacterium tuberculosis in clinical samples.
J. Clin. Microbiol. 32:672–678.
23. Ley, B. E., C. J. Linton, D. M. C. Bennett, H. Jalal, A. B. M. Foot, and M. R.
Millar. 1998. Detection of bacteraemia in patients with fever and neutrope-
nia using 16S RNA gene amplification by polymerase chain reaction. Eur.
J. Clin. Microbiol. Infect. Dis. 17:247–253.
24. Martin, A. B., S. Webber, J. Fricker, R. Jaffe, G. Demmler, D. Kearney, Y.-H.
Zhang, J. Bodurtha, B. Gelb, J. Ni, T. Bricker, and J. A. Towbin. 1994. Acute
myocarditis: rapid diagnosis by PCR in children. Circulation 90:330–339.
25. McKenzie, R., and L. G. Reimer. 1987. Effect of antimicrobials on blood
cultures in endocarditis. Diagn. Microbiol. Infect. Dis. 8:165–172.
26. Paisley, J. W., and B. A. Lauer. 1994. Pediatric blood cultures. Clin. Lab.
27. Pittet, D., D. Tarara, and R. P. Wenzel. 1994. Nosocomial bloodstream
infection in critically ill patients. Extra length of stay, extra costs, and attrib-
utable mortality. JAMA 271:1598–1601.
28. Rattanathongkom, A., R. W. Sermswan, and S. Wongratanacheewin. 1997.
Detection of Burkholderia pseudomallei in blood samples using polymerase
chain reaction. Mol. Cell. Probes 11:25–31.
29. Reimer, L. G., M. L. Wilson, and M. P. Weinstein. 1997. Update on detection
of bacteremia and fungemia. Clin. Microbiol. Rev. 10:444–465.
30. Rello, J., M. Richart, B. Mirelis, et al. 1994. Nosocomial bacteremia in a
medical-surgical intensive care unit: epidemiologic characteristics and fac-
tors influencing mortality in 111 episodes. Intensive Care Med. 20:94–98.
31. Rohner, P., B. Pepey, and R. Auckenthaler. 1997. Advantage of combining
resin with lytic Bactec blood culture media. J. Clin. Microbiol. 35:2634–2638.
32. Rudolph, K. M., A. J. Parkinson, C. M. Black, and L. W. Mayer. 1993.
Evaluation of polymerase chain reaction for diagnosis of pneumococcal
pneumonia. J. Clin. Microbiol. 31:2661–2666.
33. Smith, R. L., S. M. Meixler, and M. S. Simberkoff. 1991. Excess mortality in
critically ill patients with nosocomial bloodstream infections. Chest 100:164–
34. Smith, S. M., and R. H. K. Eng. 1983. In vitro evaluation of the Bactec
resin-containing blood culture bottle. J. Clin. Microbiol. 17:1120–1126.
35. Song, J.-H., H. Cho, M. Y. Park, D. S. Na, H. B. Moon, and C. H. Pai. 1993.
Detection of Salmonella typhi in the blood of patients with typhoid fever by
polymerase chain reaction. J. Clin. Microbiol. 31:1439–1443.
36. Sparwasser, T., T. Miethke, G. Lipford, K. Borschert, H. Ha ¨cker, K. Heeg,
and H. Wagner. 1997. Bacterial DNA causes septic shock. Nature 386:336–
37. Spencer, R. S. 1988. Blood cultures: where do we stand? J. Clin. Pathol.
38. Vincent, J.-L., D. J. Bihari, P. M. Suter, H. A. Bruining, J. White, M.-H.
Nicolas-Chanoin, M. Wolff, R. C. Spencer, and M. Hemmer. 1995. The
prevalence of nosocomial infections in intensive care units in Europe. JAMA
39. Washington, J. A. 1989. Blood cultures: an overview. Eur. J. Clin. Microbiol.
Infect. Dis. 8:803–806.
40. Weinstein, M. P. 1996. Current blood culture methods and systems: clinical
concepts and interpretation of results. Clin. Infect. Dis. 23:40–46.
41. Wilson, M. L., and M. P. Weinstein. 1994. General principles in the labora-
tory detection of bacteremia and fungemia. Clin. Lab. Med. 14:69–82.
42. Yagupsky, P., and F. S. Nolte. 1990. Quantitative aspects of septicemia. Clin.
Microbiol. Rev. 3:269–279.
43. Yamashita, Y., S. Kohno, H. Koga, K. Tomono, and M. Kaku. 1994. Detec-
tion of Bacteroides fragilis in clinical specimens by PCR. J. Clin. Microbiol.
44. Zhang, Y., D. H. Isaacman, R. M. Wadowsky, J. Rydquist-White, J. C. Post,
and G. D. Ehrlich. 1995. Detection of Streptococcus pneumoniae in whole
blood by PCR. J. Clin. Microbiol. 33:596–601.
45. Ziegler, R., I. Johnscher, P. Martus, D. Lenhardt, and H.-M. Just. 1998.
Controlled clinical laboratory comparison of two supplemented aerobic and
anaerobic media used in automated blood culture systems to detect blood-
stream infections. J. Clin. Microbiol. 36:657–661.
2482HEININGER ET AL.J. CLIN. MICROBIOL.