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Applied and Environmental Microbiology, September 1999, p. 4255-4260, Vol. 65, No. 9
0099-2240/99/$04.00+0
Bacterial Spores Survive Treatment with Commercial
Sterilants and Disinfectants
Jose-Luis
Sagripanti* and
Aylin
Bonifacino
Molecular Biology Branch (HFZ-113), Division
of Life Sciences, Office of Science and Technology, Center for Devices
and Radiological Health, Food and Drug Administration, Rockville,
Maryland
Received 22 February 1999/Accepted 9 June 1999
 |
ABSTRACT |
This study compared the activity of commercial liquid sterilants
and disinfectants on Bacillus subtilis spores deposited on three types of devices made of noncorrodible, corrodible, or polymeric material. Products like Renalin, Exspor, Wavicide-01, Cidexplus, and
cupric ascorbate were tested under conditions specified for liquid
sterilization. These products, at the shorter times indicated for
disinfection, and popular disinfectants, like Clorox, Cavicide, and
Lysol were also studied. Data obtained with a sensitive and quantitative test suggest that commercial liquid sterilants and disinfectants are less effective on contaminated surfaces than generally acknowledged.
| |
TEXT |
Different reports agree that 5 to
10% (1.75 to 3.5 million) of the 35 million patients annually admitted
to hospitals in the United States acquire an infection during
hospitalization (5, 6, 22). More than 850,000 of these have
been estimated to be implant- and device-related infections
(2). Abundant data linking the transmission of various
diseases (including AIDS, tuberculosis, and Creutzfeldt-Jakob disease,
as well as hospital epidemics of infections with
Pseudomonas, Serratia, and Bacillus species) to medical devices suggest that the effectiveness of disinfection and sterilization practices has been overestimated (21).
The capacity to kill bacterial spores determines how a commercial
product will be marketed. Disinfectants are not expected to kill all
bacterial spores and are used to decontaminate devices that ordinarily
do not penetrate tissues or that touch only intact skin (3, 16,
25). Sterilants are expected to kill all microorganisms, including bacterial spores, and are used to treat devices that penetrate tissue or present a high risk if unsterile. Viable spores still attached to various materials could remain undetected by current
sporicidal tests (1), resulting in overestimation of the
sporicidal activity of sterilizing agents (4, 7, 11, 12, 14,
15). The goal of this study was to compare the sporicidal activities of commercial liquid sterilants under manufacturer-specified conditions by using a sensitive method able to quantitatively account
for the survival of all spores on contaminated carrier devices.
Selection of carrier devices.
The device to which spores are
attached might alter the sporicidal activity of some germicidal agents
(19). Therefore, the criteria used to select the carrier
devices that we tested were based on the following practical
considerations: (i) diverse material composition, (ii) geometry
representative of medical devices, (iii) similar spore load capacities,
(iv) size amenable to microtesting, and (v) cost. Miniature stainless
steel machine screws (no. 0/80, pan head, 1.5 mm in diameter, and 12.5 mm long) were purchased at a local hardware store (Home Depot,
Rockville, Md.) or from Thompson & Cooke (Bladensburg, Md.). Dental
burs (FG 557) made of carbon steel were manufactured by Midwest Dental
Products Corporation (Des Plaines, Ill.). Medical-grade silicone rubber
tubing, 3.1-mm outer diameter and 1.5-mm inner diameter (Silastic;
catalog no. 602-285), was manufactured by Dow Corning Corporation
Medical Products (Midland, Mich.) and used in 12.5-mm-long sections.
All devices were cleaned prior to use by washing with detergent,
rinsing three times with distilled water, washing once in acetone, and rinsing again in distilled water before sterilization by autoclaving. The devices were immersed 5 mm deep in spore-loading suspensions. This
procedure contaminated areas of 20, 40, and 78 mm2 on
dental burs, screws, and tubing, respectively. Likely due to
differences in geometry and materials, the test described below loaded
similar numbers of spores onto the three devices in spite of the
different immersed areas. The miniature stainless steel screws and
small sections of medical-grade silicone rubber tubing were small
enough to fit our microtest format and inexpensive (costing 6 and 3 cents each, respectively). Easy availability of tubing, burs, and
screws made custom manufacturing of carriers unnecessary. Their low
cost allowed these carriers to be used only once and then discarded,
thus preventing spore carryover and the need to wash and sterilize the
carriers between tests.
Direct measurement of spores loaded onto carriers.
Spores of
Bacillus subtilis subsp. globigii (Spordex) were
purchased from AMSCO American Sterilizer Co. (Erie, Pa.) with a reported D value for dry-heat killing at 160°C of 2.2 min
and a D value for ethylene oxide killing (600 mg/liter at
54°C) of 3.5 min, respectively. The number of spores loaded onto
carriers was determined by using radioactively labeled spores. A method that produces dry-heat-resistant spores in synthetic medium (8, 13, 23) was modified in our laboratory so that it would result in
maximum incorporation of radiolabeled precursor as previously described
(19). A rapidly growing culture (106 bacteria in
5 ml) was inoculated into 300 ml of synthetic sporulating medium in
which methionine was replaced with radioactive
L-[methyl-14C]methionine (0.33 Ci/ml; NEC165H; 50 mCi/mmol; New England Nuclear, Boston, Mass.). After
5 days of incubation at 32°C in a shaker operating at 140 rpm,
cultures were chilled in ice and spores were pelleted by centrifugation
for 30 min at 900 × g in a Beckman TJ-R refrigerated
centrifuge. After five cycles of centrifugation and resuspension in new
Luria-Bertani (LB) broth, the radioactivity in the supernatant was
reduced to less than 2% of the radioactivity in the pellet containing
the spores. Samples from each batch of spores radioactively labeled and
concentrated in our laboratory or nonradiolabeled spores obtained
commercially (Spordex) were microscopically examined and exposed to
acid for confirmation of spore morphology and chemical resistance as
previously described (18). No vegetative cells (rods) were
observed during the counting of 1,000 radioactively labeled or
nonlabeled spores. Spores were exposed for various time periods to
either deionized, glass-distilled, autoclave-sterile water (controls)
or hydrochloric acid (2.5 N). After exposure they were neutralized with
ice-cold LB broth (Advanced Biotechnology IC, Columbia, Md.) and
titrated onto broth-agar (LB broth [Miller-Difco, Detroit, Mich.],
1.5% Agar Select [Gibco-BRL, Paisley, Scotland]) plates 100 mm in
diameter. Typical spore survival in hydrochloric acid for 5 and 10 min
was 100 and 88%, respectively.
Spores labeled with [14C]methionine were diluted in LB
broth, and identical aliquots were either titrated for viability or
counted for radioactivity. The specific activity of each spore
preparation was obtained from the slope of the regression line of spore
number (as determined by titration) versus incorporated 14C
label (measured by scintillation counting). We transferred various devices to Eppendorf polypropylene tubes (1.5 ml) containing 50 µl of
14C-labeled spores at different concentrations. Each device
was immersed in a separate spore-loading suspension for 30 min. The devices were then removed from the loading suspension with forceps and
dried for 10 min under vacuum (Speed Vac; Savant, Farmingdale, N.Y.).
Each 50-µl suspension was used once and then discharged.
The spore load on each device was estimated by immersing the loaded
devices in scintillation liquid, measuring radioactivity,
and
multiplying this value by the specific activity of the preparation.
One
large batch with a specific activity of 1.7 × 10
3 ± 0.3 × 10
3 spores per cpm was
used for final calibration of all devices.
The number of spores
attached to no. 0/80 stainless steel screws
(ranging from 6.0 × 10
6 to 6.5 × 10
6) was comparable to that
loaded into medical-grade silicone rubber
tubing (3.8 × 10
6) immersed in a spore suspension with a similar spore
concentration
(1.7 × 10
9/ml). The increase in the
number of spores loaded onto the stainless
steel screws or silicone
rubber tubing was approximately linear
with increasing concentrations
of the loading suspension in the
range of 10
7 to
10
10 spores/ml. This contaminating procedure loaded, on
average, 3
spores per 1,000 spores/ml of the loading
suspension.
Sterilants and disinfectants.
Cidexplus (3.4% glutaraldehyde,
pH 8.0; Johnson and Johnson Medical Inc., Arlington, Tex.) was
activated as specified and used full strength at 21°C over a period
of either 10 h, for sterilization, or 20 min, as indicated for
high-level disinfection. Exspor (Alcide Corp., Redmond, Wash.),
containing 1.52% sodium chlorite, was activated daily before
experiments by mixing 1 part base concentrate, 4 parts water, and 1 part activator (yielding a pH between 2.3 and 2.7). The label
prescribes the treatment of medical items with an Exspor-activated
solution for 10 h to achieve sterilization and for 1 to 3 min for
killing of Mycobacterium sp. and other bacteria, pathogenic
fungi, and viruses on hard surfaces. Renalin (Renal Systems Division of
Minntech Corp., Minneapolis, Minn.), a mixture of 20.0% hydrogen
peroxide and 4.0% peroxyacetic acid, was used as recommended for
sterilization at a dilution of 1:5 (final dilution; pH 1.8) in sterile,
deionized, and glass-distilled water for an 11-h exposure. Wavicide-01
(2% glutaraldehyde; Wave Energy Systems, Wayne, N.J.) was used full
strength for 10 h at 21°C as a sterilant or at a 1:4 dilution
for 10 min (at room temperature [21°C]), as specified for killing
of vegetative bacteria and viruses. Clorox (5.25% sodium hypochlorite,
manufactured by The Clorox Company, Oakland, Calif.) was used at a 1:21
dilution, as recommended for disinfection. Lysol I.C. (7.24%
o-benzyl-p-chlorophenol and 2.23%
o-phenylphenol; National Laboratories, Montvale, N.J.) was used at the 1:128 dilution specified for use in hospitals, nursing homes, dental offices, and other institutional facilities as a germicidal, tuberculocidal, pseudomonacidal, staphylococcidal, fungicidal, and virucidal compound. Cavicide (15.30% isopropanol and
0.25% diisobutyl phenoxyethoxyethyl dimethyl benzyl ammonium chloride;
Micro Aseptic Products, Inc., Palatine, Ill.) was used full strength,
as specified for disinfection of noncritical medical instruments.
Cupric chloride (CuCl2 · 2H2O;
Mallinckrodt Specialty Chemicals, Paris, Ky.), L-ascorbic
acid, and (30% wt/vol) hydrogen peroxide (both from Aldrich Chemical,
Milwaukee, Wis.) were used in a mixture (0.5% cupric ions [as cupric
chloride]-0.1% ascorbic acid-0.003% hydrogen peroxide, pH 2.9).
Sporicidal test on contaminated medical devices.
Each carrier
device was independently immersed in a tube with 50 µl of a
suspension of radiolabeled spores (1.7 × 109
spores/ml). After drying, the devices were divided into two identical groups. In one group, the number of spores loaded into each device was
measured radioactively. Devices in the second group were incubated at
20°C in 400 µl of disinfectant (three devices per disinfectant in
separate tubes) for the time period specified on the respective product
label or in an equal volume of sterile distilled water for 30 min, as a
control for spore survival. After incubation, each device was removed
from the test tube, the remaining disinfectant was diluted with 600 µl of ice-chilled LB broth, and the tube was centrifuged (5 min at
15,000 rpm in a model 5414 Microfuge [Brinkman Instruments Inc.,
Westbury, N.Y.]). The supernatant with diluted disinfectant was
discarded; the spores in the pellet were resuspended by vortexing in
fresh, ice-chilled LB broth (1 ml); and this sample containing loosely
adherent spores was named fraction a. The device removed in the step
described above was transferred to 400 µl of distilled water and
sonicated for 5 min (Ultrasonic Cleaner; Cole Parmer, Chicago, Ill.).
After sonication, the device was removed and 600 µl of ice-chilled LB
broth was added to the 400 µl of water. This sample with spores
removed by sonication was named fraction b. To recover viable spores
still remaining on the carriers after fractions a and b had been
obtained, the devices were incubated in 400 µl of fresh LB broth for
30 min at 32°C in a shaker operating at 140 rpm. The device was
removed and counted in scintillation liquid, and lack of radioactivity confirmed the absence of detectable spores. Six hundred microliters of
ice-chilled LB broth was added to the broth left after device removal,
and this sample with spores dislodged after 30 min of shaking in medium
was named fraction c. The incubation time of fraction c (30 min) was
shorter than the period required for spores of B. subtilis
to germinate and replicate, thus preventing overestimation of surviving
organisms (data not shown). Fractions a, b, and c were serially diluted
in LB broth, and the surviving spores in each fraction were titrated by
serial dilution on LB broth agar plates.
The overall recovery ratio of the method was calculated as the sum of
spores titrated in fractions a, b, and c (ranging from
2.9 × 10
6 to 10.9 × 10
6 spores) after treatment
with water divided by the average number
of spores loaded (estimated
radioactively). The spore recovery
of the three-step method was
1.02 ± 0.22 (average fraction of
the starting spore number ± the standard error (SE) in six independent
experiments) for
0/80 stainless steel screws, a value nearly identical
to the recovery
previously obtained for silicone catheter tubing
(1.02 ± 0.59).
The recoveries of nonradiolabeled or radiolabeled
spores in fractions
a, b, and c were similar with all of the devices
studied. Therefore,
nonradioactive spores were used after the
number of spores loaded onto
each device was calibrated and it
was established that the three-step
method accounted for all of
the challenge spores. By using the same
devices and procedure,
other laboratories could reproduce this test
without further calibration
or need for radioactive
spores.
We included positive and negative controls for sporicidal activity in
each experiment to allow monitoring of intertest performance.
Water was
chosen as the negative control because of its lack of
sporicidal
activity and common availability (no killing or 100%
spore survival).
Stability in dry chemical form and relatively
low cost made cupric
ascorbate a convenient positive control for
sporicidal activity that
produced a significant, consistent, and
relatively time-independent
(between 30 min and 10 h) reduction
in spore survival (see Table
1).
The sporicidal test that we developed has several valuable
characteristics. (i) It is quantitative. The number of spores attached
to the devices before disinfection was directly measured with
radiolabeled spores. Absence of spore attachment to the carrier
at the
end of the testing process is easily confirmed by determining
lack of
remaining radioactivity. A recovery value of nearly 1
in the negative
controls demonstrates that all of the loaded spores
are accountable for
by the test. This is a clear advantage over
methods that estimate
carrier load indirectly by measuring the
spores dislodged from the
device to an unknown extent. Furthermore,
determining the surviving
fraction at each step of the test by
counting colonies from surviving
spores is more precise and informative
than scoring growth or nongrowth
as in other sporicidal tests.
(ii) It is rapid. Our procedure was
completed within 4 h, not
counting overnight colony development.
(iii) It is economical
and environmentally friendly. The technique uses
only 400 µl of
disinfectant, resulting, for all practical purposes,
in a nondestructive
test that saves reagents and reduces the amount of
toxic and infectious
waste
produced.
Effect of germicides on contaminated devices.
Devices carrying
3.8 × 106 to 6.2 × 106 spores of
B. subtilis were exposed once to various sterilizing agents
or to water, and the spores titrated in fractions a, b, and c are shown
in Fig. 1 (tubing) and
2 (screws). It was unclear how much the
sporicidal activity of products labeled as liquid sterilants differed
from that of common disinfectants. To answer this question, we also measured the relative sporicidal activities of products not intended for liquid sterilization but recommended for disinfection of medical devices used in patients with AIDS or decontamination of surfaces during epidemics or bacteriological warfare or widely used as household
disinfectants (9, 10, 20, 24). The spore survival results
shown in Fig. 1 and 2 and Table 1 confirm
that general disinfectants (not specifically labeled for liquid
sterilization, like Cavicide, Clorox, and Lysol) do not kill spores on
contaminated devices and, thus, should never be employed in this
capacity.
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FIG. 1.
Effects of germicides on spores deposited on silicone
rubber medical tubing. Spores were loaded onto tubing, dried, and
exposed for the times prescribed on the products' labels either for
sterilization or for disinfection (indicated on the X axis).
Viable spores were measured in fractions a, b, and c as described in
the text. Bar height represents mean survival, and the bracket over the
bars indicates the SE of triplicate determinations in three to eight
independent experiments.
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FIG. 2.
Spore survival after treatment of stainless steel
screws. Reagents and conditions were as described in the text and in
the legend to Fig. 1. Bar height represents the mean ± the SE of
the number of viable spores obtained in fractions a, b, and c
determined in triplicate by titration in four to eight independent
experiments.
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|
Figures
1 and
2 show that the proportion of viable spores recovered in
each fraction varies for different products and treatment
times. For
all products, a one-step procedure (fraction a with
loosely adherent
spores) failed to detect all of the spores remaining
viable after
treatment (Fig.
1 and
2). Spore recovery in fraction
a was often lower
than after sonication and 30 min of shaking
in culture medium,
respectively (fractions b and c in Fig.
1 and
2). This could be a
consequence of fixing or trapping of viable
spores on the device
surface by chemical cross-linking with the
germicide. No viable spore
could be detected in fraction a after
treatment of tubing with the most
active disinfectant (Renalin
incubated for 11 h; Fig.
1). However,
by using a procedure that
completely recovers attached spores, a few
spores were detected
after sonication in fraction b and several hundred
spores were
easily detected after 30 min of shaking in medium (fraction
c).
Often, the number of surviving spores detected in fraction a
differed
by more than 1 log from the total number of viable spores
(fractions
a, b, and c in Fig.
1 and
2). Products whose effectiveness
would
be overestimated by more than 10-fold by a one-step recovery
method
included Cavicide, Cidexplus, Exspor, Lysol, and Renalin. Thus,
these findings indicate that the sporicidal activity of disinfectants
and sterilants will likely be overestimated by methods that dislodge
spores in only one step (obtaining results equivalent to those
obtained
with fraction a) or by tests in which the recovery of
loaded spores is
unknown.
Comparative sporicidal activity.
The total log of spore
killing was obtained by subtracting the log of the total number of
viable spores after exposure of devices to disinfectants (titrated in
fractions a, b, and c) from the log of the number of spores surviving
treatment with water. The values obtained for each device-disinfectant
combination are displayed in Table 1. The survival of spores on
contaminated dental burs was higher than on the other two devices.
Disinfection of carbon steel dental burs produced corrosion stains on
the devices and a fine precipitate at the bottoms of the test tubes.
The higher spore survival correlated with obvious corrosion, and
therefore, data on burs were not considered for comparison or ranking
of products. The severe corrosion observed after treatment with
commercial disinfectants made carbon steel dental burs inadequate as
carriers for sporicidal testing. Deterioration after a single test and increased spore survival demonstrate that dental burs (and likely other
devices containing carbon steel) must not be decontaminated with liquid
disinfectants in spite of instructions to the contrary on the labels of
some carbon steel devices. In contrast, the other two materials in this
study were impervious to all disinfecting treatments. Stainless steel
screws and silicone rubber catheter tubing did not show signs of
deterioration after visual and microscopic examination (×160
magnification). These findings agree with the relative resistance of
stainless steel and medical-grade silicone rubber to corrosion
(17). Similar spore recovery and killing (within 1 log) by
the same disinfectant on both devices (Table 1) suggest that testing on
stainless steel screws and medical silicone rubber tubing should
provide an adequate estimation of sporicidal activity on medical devices.
Cidexplus is specified to be used for up to 28 days after activation.
The label of Renalin indicates that the diluted solution
must be used
within a 7-day period as a sterilant for dialyzer
reprocessing. These
sterilants were tested at various times after
activation or dilution.
No significant change in the sporicidal
activity of Cidexplus or
Renalin was detected on contaminated
silicone tubing, dental burs, and
stainless steel screws during
a 28- or 7-day test period, respectively
(data not
shown).
The incubation time specified in the labeling of products intended for
sterilization is 10 or 11 h. Much shorter incubation
times (a few
minutes) are specified for use of the same products
as sterilants.
Changes in incubation time had a distinct effect
on spore killing
produced by different formulations specified
as sterilants (Table
1).
Extending treatment with Wavicide-01
from 10 min to 10 h caused a
relatively large increase (more than
100 times) in sporicidal activity.
In contrast, extending treatment
with Exspor or cupric ascorbate from a
few minutes to 10 h did
not produce a substantial increase in
sporicidal activity (less
than a 10-fold difference between short and
long exposures). Unexpectedly,
spore killing on screws was slightly
higher after 20 min than
after 10 h of incubation with Cidexplus
in four independent comparative
experiments (Table
1). These findings
suggest that the sporicidal
activity of some products may be exhausted
after a relatively
short incubation period and highlight the importance
of precise
adherence to the times specified by the particular
product's
label.
Glutaraldehyde and peroxi compounds are common active ingredients used
in liquid sterilization and high-level disinfection
(
3,
9,
10,
16,
21,
25). However, commercial products
with these active
ingredients had quite different sporicidal potencies
after incubation
for the similar periods (10 and 11 h of treatment)
recommended for
sterilization. The reduction of spore numbers
ranged from 2,500- to
56,000-fold for Cidexplus and Renalin, respectively
(Table
1).
The substantial spore survival detected in this study after treatment
of devices with commercial sterilants (Table
1) conflicts
with the
concept of sterilization, defined as the destruction
of all life,
including bacterial spores. The data that we obtained
with a sensitive
and quantitative test suggest that commercial
liquid sterilants and
disinfectants are less active on contaminated
surfaces than generally
acknowledged.
| |
ACKNOWLEDGMENTS |
We appreciate the critical review of our manuscript by Larry E. Bockstahler (Division of Life Sciences, CDRH, FDA, Rockville, Md.),
review of the statistical analysis by Harry F. Bushar (Division Of
Biostatistics, CDRH, FDA, Gaithersburg, Md.), and assistance in
measuring the contaminated areas of devices by Robert Bolster (Naval
Research Laboratory, Washington, D.C.).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Molecular
Biology Branch (HFZ-113), Center for Devices and Radiological Health,
Food and Drug Administration, 5600 Fishers Ln., Rockville, MD 20857. E-mail: JUS{at}CDRH.FDA.GOV.
| |
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