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The sterility assurance level of 10−6 is an established standard that defines the quality of sterile products. The aim of the present study was to develop a method that correlated the results from microbial-barrier testing of flexible sterile barrier systems with the estimated microbial challenge that the package encounters during storage and transport.
The effectiveness of microbial-barrier packaging was determined by the use of an exposure chamber test with 20 periodic atmospheric pressure changes of 50 and 70 hPa. Flexible peel pouches were used as sterile barrier systems. The logarithmic reduction value of a sterile barrier system was calculated on the basis of the experimental results and compared with the logarithmic reduction value required for the microbial challenges to maintain sterility during transport and storage.
For pouches made of paper and plastic-film material, a logarithmic reduction value of 5.4 was obtained on the basis of 30 of 99 plates becoming nonsterile after being exposed to a 50 hPa difference in periodic atmospheric pressure changes. For pouches made of paper and plastic-film material, a logarithmic reduction value of 5.2 was obtained on the basis of 48 of 100 plates becoming nonsterile after being exposed to a 70 hPa difference in atmospheric pressure. For pouches made of nonwoven and plastic-film material, logarithmic reduction values of 6.38 (ie, 3 of 99 plates became nonsterile after being exposed to a 50 hPa pressure difference) and 6.07 (ie, 3 of the 99 plates became nonsterile after being exposed to a 70 hPa pressure difference) were obtained. Calculating an expected microbial challenge during transport and storage that requires barrier properties corresponding to a logarithmic reduction value of 5.83 and taking the sterility assurance level into account, we found that only the nonwoven pouches fulfilled the European standard EN 556-1.
Using the data obtained in a microbial exposure test with a specified flow rate of a bacterial aerosol, we found that the effectiveness of the sterile barrier system against the actual microbial challenge can be examined and evaluated at the sterility assurance level of 10−6.
We developed a microbiological test to detect the penetration of airborne microorganisms through the packaging of medical products after sterilization, to meet the requirements of European standard EN 556. We applied this test method to transparent pouches.
The microbial-barrier properties of the transparent pouches were determined using the microbial challenge test, in which the package was placed inside an exposure chamber and exposed to a defined aerosol of Saccharomyces cerevisiae. The atmospheric pressure in the chamber was periodically reduced by 0-75 millibars, to simulate weather-dependent pressure changes. Thermoresistant petri dishes filled with nutrient agar were integrated into the transparent pouches before sterilization. The packages were incubated after exposure. They were then opened and examined for colony growth.
The number of recontaminated packages per test group (n = 50) depended on the microbial bioload (defined as the number of colony-forming units per plate) to which the packages were exposed and on the size and number of decreases in atmospheric pressure. Results of multiple regression analysis showed a significant increase in the number of recontaminated packages in correlation with the product of the values for microbial bioload and the size and number of decreases in atmospheric pressure. When we analyzed the probability of recontamination of wrapped medical devices after 2 reductions in atmospheric pressure (30 millibars each) and with a surface microbial load of 10 colony-forming units per 64 cm2, we estimated that the frequency of recontamination was 1: 100,000.
Multiple regression analysis showed that the proposed microbial challenge test is suitable to determine the probability of package recontamination at the 1: 1,000,000 level.
To develop a microbial test method to ascertain the passage of airborne bacteria through the medical device packaging System after sterilization, and to apply this test method to flexible packages under mechanical pressure changes.
Petri dishes filled with nutrient agar were integrated into the packaging unit prior to sterilization. We examined paper packaging consisting of 1 (single-paper packaging [P]), 2 (double-paper packaging [PP] and textile and paper double packaging [TP]), and 3 (double packaging with transport packaging [TPP]) layers. After sterilization, the test packages were pressed five times per minute for 1 or 3 hours by a mechanical device weighing 1 kg. This exposure took place in rooms with an average airborne bacterial count of 35 (room 1) or 440 (room 2) CFU/m3. The packaging was opened following culture at 37° C for 48 hours to determine the number of colonies formed.
The proportion of contaminated packages rose with the duration of mechanical stress and increased airborne bacteria concentration. Thus, mechanical pressure change for 3 hours resulted in the contamination of 60% (P), 15% (PP), 0% (TP), and 0% (TPP) of the packages in room 1, whereas 100% (P), 65% (PP), 73% (TP), and 0% (TPP) of the packages in room 2 were contaminated.
This test method allows sterile packaging Systems to be tested for contamination under practical conditions without extensive laboratory preparation. Contamination as a resuit of laboratory errors can be ruled out almost certainly.
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