Key words:
Microbial contamination - paper/plastic pouches - resterilization - sterility - storage
time
INTRODUCTION
Steam sterilization of medical/dental equipment is an important process to prevent
infection in patients undergoing medical/dental procedures.[1] One critical aspect of sterilization is packaging, which plays an important role
in preserving the sterility of medical/dental equipment and in preventing microbial
contamination from the external environment after the sterilization process.[2] Paper/plastic peel pouches are widely used packaging materials for steam sterilization
with an autoclave due to its convenience of use, content visibility, and efficacy.[3] The paper side is composed of complex cellulose fibers, and the other side is made
of laminated transparent polyester/polypropylene.[3]
[4] Each side is fused together by heat sealing.[4] The pouches should be inspected to ensure complete closure of the package before
and after sterilization.
Following the sterilization process, the equipment is generally not used immediately
but stored for later use. Safe storage depends on the conditions of the storage environment.
Previous studies found that storing sterilized equipment in open environments, such
as open shelves, resulted in faster microbial penetration than storing in closed cabinets
with dustcovers,[5] and that using different packaging materials (reusable woven packs, disposable nonwoven
packs, and polypropylene peel pouches) had no effect on safe storage time.[6] This indicates that the storage environment may be a more important factor than
the type of packaging material in maintaining sterility.
A survey on autoclave dental packaging in Thailand found that paper/plastic pouch
was the most commonly used packaging material for steam sterilization in both hospitals
and private clinics.[7] Although these pouches are recommended for single use by the manufacturer,[8]
[9] almost all of these clinics reused the pouches. The most frequent times of resterilization
were 3 times with 5 as the maximum times of reuse. However, the impact of pouch reuse
and the integrity of reused pouches were not explored in that survey. Indeed, the
work exploring the effects of resterilization on the packaging material is scarce.
Most studies regarding resterilization focus on medical/dental instruments themselves,
not on the packaging.[10]
[11] Even though packaging material is also an important factor in sterility maintenance.
Various factors can lead to loss of sterility of these pouches including the barrier
efficacy of the paper/plastic pouch itself, packaging process, environmental factors
(closed or open), and mishandling of the packages from human error. One critical factor
of pouch reuse is the deterioration of barrier efficacy of these paper/plastic pouches
after repeated sterilization processes. If resterilization process impaired the sterile
integrity of the pouches and resulted in shorter storage time or risk of contamination,
this information would be useful when considering reusing these pouches. The objective
of this study was to determine the barrier efficacy of paper/plastic pouches on the
sterility maintenance after resterilizing 1, 3, and 5 times and after storage in a
closed environment for up to 6 months.
MATERIALS AND METHODS
Study protocol and preparation of paper/plastic pouches
There were four experimental groups and two control groups as shown in [Figure 1].
Figure 1: Details of the experimental and control groups
The paper/plastic pouches were prepared from sterilization tubular rolls (Stericlin
see Through Reels, VP Medical Packaging, Germany) at 7.62 cm × 12 cm apiece. Each
pouch contained a piece of 0.5 cm × 2 cm filter paper (Whatman paper, Patterson Scientific,
England) and a piece of internal chemical indicator (3M Comply SteriGage Chemical
Integrator, 3M, USA) inside. Then, the pouches were sealed 1 cm from the bottom and
3 cm from the top with a heat sealer (Euroseal, Euronda S.p.A., Italy) as shown in
[Figure 2a]. Each sealed pouch was inspected for bubbles, gaps, folds or creases, and holes
and burn through to ensure the integrity. Resterilized pouches were not sealed in
the first cycles; they were sealed and put in the filter paper and internal chemical
indicator only in the last cycle of sterilization [Figure 2b]. In the positive control group, the pouch was punched through both paper and plastic
sides, and two cuts were also made at the sides of the pouch to deteriorate barrier
integrity [Figure 2c]. Autoclave tape (3M ESPE Autoclave Steam Indicator Tape, 3M, USA) was placed on
the plastic side of the pouch. The pouches were arranged in vertical position and
the paper side was in contact with the plastic side of the next pouch without touching
the chamber wall of an autoclave (M11 UltraClave, Midmark Corporation, USA). All samples
were sterilized at 121°C and 15 psi for 30 min.
Figure 2: Paper/plastic pouch preparation. (a) Sealed pouch containing filter paper and internal
chemical indicator. (b) Unsealed pouch sterilized in earlier cycles for R1, R3, and
R5 groups. (c) Positive control
Sterilization monitoring
Three modes of monitoring were applied to every sterilization cycle: physical, chemical,
and biological. Physical monitoring was direct observation of the gauges on the autoclave
machine during the sterilization process. Chemical monitoring was done using an internal
chemical indicator and autoclave tape as an external chemical indicator. Both types
of chemical indicators would change the color following sterilization process. For
biological monitoring, spore test tubes (3M Attest, 3M, USA) were placed at the center
and opposite corners of the autoclave tray. After the sterilization process, the spore
test tubes were incubated at 56°C for 48 h to evaluate for microbial growth.
Storage
All sterilized samples were stored in closed plastic boxes for 1, 2, 3, 4, 5, or 6
months. Each box contained 125 samples (30 from each experimental group and 5 from
the positive control group).
Microbial cultivation
After the specified storage period, the pouch was inspected for barrier damage before
opening. The filter paper was aseptically removed and incubated in Brain Heart Infusion
(BHI) broth (Becton Dickinson, Maryland, USA) at 37°C for up to 2 weeks. The turbidity
of the media indicated microbial contamination.
RESULTS
Physical monitoring during the sterilization process showed that the conditions required
were met, i.e., the temperature reached 121°C and the pressure was at 15 psi in every
cycle of sterilization. Both internal pouch with holes and cuts to impair barrier
integrity and external chemical indicators were inspected on every package and all
showed color change indicating the proper functioning of the autoclave machine. Biological
monitoring with the spore test also showed negative growth in all samples indicating
sterile status inside the autoclave.
All filter paper retrieved from new pouches, and 1 time, 3 times, and 5 times resterilized
pouches demonstrated no microbial contamination after storage in a closed environment
for up to 6 months [Table 1]. The negative control group was a new pouch sterilized on the same day that the
microbial culture of the positive control group and experimental groups was carried
out, without any storage. It represented the gold standard of pouch integrity. All
pouches in the negative control group showed no microbial contamination. All samples
in the positive control group (intentionally damaged pouches) showed microbial contamination
at every storage period. The presence of positive microbial growth was observed mostly
(90.3% of all positive control samples) within 24 h after incubation and all samples
showed positive results within 8 days of culture [Figure 3].
Table 1:
Results on microbial culture from new, resterilized, and control groups
|
Storage time (month)
|
Experimental groups
|
Control groups
|
|
N
|
R1
|
R3
|
R5
|
Negative control
|
Positive control
|
|
+
|
−
|
+
|
−
|
+
|
−
|
+
|
−
|
+
|
−
|
+
|
−
|
|
*Same condition with new pouch (N) at this time point. +: Positive microbial culture,
−:Negative microbial culture
|
|
0
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
-*
|
-*
|
40
|
0
|
|
1
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
40
|
0
|
|
2
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
40
|
0
|
|
3
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
40
|
0
|
|
4
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
40
|
0
|
|
5
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
40
|
0
|
|
6
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
0
|
240
|
40
|
0
|
|
Total
|
0
|
1680
|
0
|
1680
|
0
|
1680
|
0
|
1680
|
0
|
1440
|
280
|
0
|
Figure 3: Time distribution for positive microbial culture of positive control group (n = 280)
DISCUSSION
Resterilized paper/plastic pouches could maintain sterility after at least up to 6
times of steam sterilization and up to 6 months of storage in a closed environment.
This indicates that repeated sterilization is not a contributing factor in deteriorating
the barrier efficacy of the paper/plastic pouches. To reuse a pouch, the used pouch
has to be inspected thoroughly on both the paper and plastic sides, as well as along
the sealing margins, for damages, tears, or holes. Areas of broken seal from peel
opening or any damages are cut out, and resealing is done at a new undamaged position.
In practice, only an undamaged pouch with adequate space for packaging and resealing
at a new position can be resterilized; thus, the pouch will be smaller after each
cycle of sterilization. However, this study was designed to investigate the barrier
efficacy of the paper/plastic pouches in maintaining sterility after repeated sterilization
in an ideal condition. Thus, filter paper was applied in place of medical/dental equipment
to omit the possibility of sharp objects damaging the pouch integrity.
A previous study on paper/plastic pouch resterilization by Palananthana et al. used stainless steel wires packed inside the pouches and determined their sterility
after being resterilized 1–5 times and stored in a closed cabinet for 4 months.[12] The results found 1.33% contamination in 1 time resterilized group at 2 months storage
time, but there was no contamination in all other experimental groups (2, 3, 4, or
5 resterilization cycles and 3–4 months in storage), as well as the new pouch group.
However, there was no negative control group to compare the findings. The results
were ambiguous since the pouches that showed contamination were the ones with fewer
cycles of sterilization (1 time resterilization) and a shorter storage time (2 months),
while the pouches with more resterilization cycles (5 times resterilization) and longer
storage time (4 months) could still maintain sterility. Our study found microbial
contamination in all samples from the positive control group. All pouches in the positive
control group were handled in a similar manner to pouches from the experimental groups
and with gloves worn during transfer. This indicates that contamination may be from
the ambient environment alone and that the barrier integrity is a very important factor
in maintaining sterility of the package.
Studies on the shelf life of new paper/plastic pouches have found contamination rates
between 0% and 1.6%.[6]
[13]
[15] With low contamination rates, we calculated the sample size as 6720, which was significantly
more than in past studies and with negative and positive controls to ensure enough
chances for potential contamination. In all studies where contamination was detected,
it was not time related but rather event related, as the negative control (without
storage) also showed contamination not significantly different from the groups with
longer storage times (up to 1 year).[13] There was the absence of contamination in the negative control group in our study.
In studies without a negative control, microbial contamination was also not associated
with storage time.[6] It was suggested that inadvertent contamination might have occurred during unpackaging
and transfer of the medical/dental equipment.[14] Similarly, a prospective study which periodically checked for sterilized items shelved
in hospital wards found no contamination for up to 2 years.[16] Indeed, with proper storage and handling conditions, the shelf life of sterilized
equipment could be longer but no study has explored beyond a 2 year period. From these
results, it can be concluded that microbial contamination is not dependent on storage
time or the number of sterilization cycles.
This study used BHI broth for microbial culture to examine potential contamination.
This medium has basically similar ingredients to Trypticase soy broth employed in
other studies.[5]
[13]
[16] It can be used to culture both aerobic and facultative anaerobic bacteria.[17] We monitored the culture results for up to 2 weeks which was similar to the time
period utilized in previous studies, as positive cultures might still be observed
in the 2nd week.[13] Most positive microbial growth occurred within 24 h, similar to the time frame suggested
by the manual,[17] and all samples were found positive within 8 days of culture. BHI appeared to be
appropriate for culture of microbial contamination in this case, as the culture time
was less than those using Trypticase soy broth, and all positive control samples consistently
showed valid microbial growth.
Resterilization conditions of paper/plastic pouches in this study imitated an ideal
condition where the sterilization process was repeated immediately with minimum handling.
In real practice, reused pouches have a higher risk of event related damage to the
sterility of the pouches from unpackaging, re packaging, handling, and transferring
of pouches and equipment, as well as human error. This study was able to reveal only
the intact barrier efficacy of these pouches after repeated sterilization. A study
on actual reuse practice with pouches containing medical/dental equipment is needed
to validate these findings before the safety of reused pouches can be confirmed.
CONCLUSIONS
Resterilization of paper/plastic pouches for up to 5 times and storage in a closed
environment for up to 6 months did not impair the barrier integrity of the pouches.
Financial support and sponsorship
This study was financially supported by Faculty of Dentistry, Mahidol University,
Bangkok, Thailand.
