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Pharmaceutical Technology, Bioprocessing Supplement 2012 (PDF)

A relatively recent advance in biopharmaceutical processing has been the use of single-use disposable technologies. Single-use disposable technologies include tubing, capsule filters, single-use ion exchange membrane chromatography devices, single-use mixers, bioreactors, product holding sterile bags in place of stainless steel vessels (i.e., sterile fluid containment bags), connection devices, and sampling receptacles (1). The drivers for the implementation of such technology include improved sterility assurance, greater flexibility for biopharmaceutical manufacturing processes, reduced processing times, and cost and energy savings associated with the avoidance of washing, drying, and sterilizing stainless-steel equipment (2). 

These advantages notwithstanding, the application of single-use sterile disposable technology in the biopharmaceutical industry remains in its infancy and there are several important validation steps to be considered. These steps relate to product compatibility issues (e.g., extractables and leachable) and to sterility assurance. This paper addresses an important aspect of sterility assurance: ensuring that single-use systems are sterile and that the process of rendering single-use systems sterile does not damage the device or cause adulteration of the product. 

The primary method for the sterilization of single-use technology is by gamma irradiation. This is because plastics cannot be subjected to heat-based methods of sterilization without damaging the mould (i.e., styrene and other plastics are temperature sensitive) (3). Other alternative methods, such as gaseous sterilization by ethylene oxide, that although used on occasions, can leave unwanted and potentially toxic residuals (e.g., ethylene glycol and ethylene chlorhydrin) post-sterilization (4). Other alternative sterilization methods (e.g., liquid peracetic acid immersion system and plasma sterilization processes) are not sufficiently well established (5). Electronic beam irradiation is an alternative method of radiation to gamma. This is a concentrated, highly-charged stream of electrons generated by the accelera tion and conversion of electricity (6). Electronic beam radiation is not commonly used for single-use disposable systems due to its relatively limited ability to penetrate some types of plastics (7). Therefore, single-use systems are typically sterilized using gamma rays (i.e., electromagnetic wave radiation) (8). Despite the widespread application of the gamma irradiation, the process remains the only primary sterilization method not described in either the 

This paper outlines the process of gamma radiation and describes the important aspects of validation. The paper is designed to provide a guide to those wishing to understand more about the process and to offer advice for quality assurance personnel who are required to audit the sterility assurance of gamma radiation. 

Single-use disposable technologies have been used in biopharmaceutical processing since the 1970s on a limited scale. It is only since the 21st century that such technologies are being implemented for large-scale manufacturing (e.g., upstream, downstream, and product filling applications). The forces driving the wider adoption include improving or eliminating cross contamination, sterility assurance, process efficiencies, operator protection, and cost savings.  

Single-use disposable technologies are generally manufactured from plastic polymers involving processes of injection molding, extruding, and blow molding. The applications include devices for making aseptic connections, sampling devices, mixing devices, product-hold bags, and disposable manifold systems. Each of these systems is used to process or contain fluids including additives, buffers, bulk intermediates, and final formulations. 

Before single-use technologies can be adopted, a number of tests must be performed in relation to design, microbial inhibition, chemical computability, and leachables and extractables (9). 

The use of ionizing radiation dates back to the discovery of X-rays by Roentgen in 1895, and gamma radiation has been an established sterilization method since the early 1970s (10). It has only been since the 21

  century, however, that the use of gamma irradiation has increased within the biopharmaceutical industry, replacing more established and costly methods like ethylene oxide. This arises from the use of gamma radiation to sterilize consumables and single-use technologies used for aseptic filling operations (11). It is the recent impetus for new cleanroom technologies that is leading to an increased use of gamma radiation as the optimal sterilization method.  

The process of gamma radiation is a form of ionization (i.e., electron disruption). Gamma radiation is one of the three types of natural radioactivity, the other two being alpha and beta radiation (12). Gamma radiation is in the form of electromagnetic rays, like X-rays or ultraviolet light, of a short (less than one-tenth of a nanometer), and thus energetic, wavelength. It provides a physical means of sterilization or decontamination as the rays pass through the product being sterilized (i.e., irradiated) (13). In doing so, the radiation kills bacteria, where there is sufficient energy, at the molecular level by breaking down bacterial DNA and inhibiting bacterial division (14).  

The most common source of gamma rays for radiation processing comes from the radioactive isotope Cobalt 60, although other radionuclides can be used, such as Cesium 137. Each element decays at a specific rate and gives off energy in the form of gamma rays and other particles. Cobalt 60 is manufactured specifically for the gamma radiation process from non-radioactive Cobalt 59. The radioactive Cobalt 60 functions as the isotope source. High-energy photons are emitted from the Cobalt 60 to produce ionization (electron disruptions) throughout a product. The gamma process does not create residuals or impart radioactivity in processed products (15). 

Gamma radiation is often referred to as a "cold process" for the temperature of the processed material that does not significantly increase (16). The sterilization process is not dependant on humidity, temperature, vacuum, or pressure, which means that the process is suitable for materials that cannot be subjected to high-temperature sterilization. The important variables for gamma radiation are the strength of the radiation dose (i.e., measurement of how much energy is absorbed when something is exposed to the radiation source) and the exposure time. The measurement of radiation is expressed in units called KiloGrays (KGy). One gray is the absorption of one joule of radiation energy by one kilogram of matter (17).  

 The regulatory requirements for gamma radiation are less defined than for sterilization by filtration, moist or dry heat, or by ethylene oxide. These methods of sterilization normally involve a direct biological challenge, as with biological indicators (preparations of a specific microorganism, with high resistance towards particular sterilization methods) for steam sterilization or a high population microbial challenge for filter validation. In the past Bacillus pumilus was used as a biological indicator to measure gamma irradiation. It was removed because the assessment of the product bioburden during validation was deemed to be a more accurate means of assessing potential resistance to the gamma radiation process. In contrast, with gamma radiation, the biological assessment is normally undertaken by an assessment of the product bioburden (i.e., the number of microorganisms on a certain amount of material prior to that material being sterilized) by assessing the total viable count (TVC) of the material, expressed as colony forming units (CFU). 

For gamma radiation, the applicable standard is ISO 11137 "Sterilization of Health Care Products-Radiation" (18). The standard was developed in association with the Association for the Advancement of Medical Instrumentation (AAMI). 

The standard is divided into three parts. The first part deals with validation and routine control methods. The second part deals with the establishment of radiation doses for items to be sterilized, and the third part relates to dosiometry. The standards function to determine how much radiation is permitted in order to achieve the desired level of sterilization when measured in terms of sterility assurance. The sterility assurance level is normally 106 (that is a theoretical concept where it is assumed that, in terms of probability, no more than one item sterilized out of one million would contain one or more microorganisms after the completion of the sterilization process) (19). 

The official scope of the ISO 11137 standards is limited to medical devices. However, in the absence of any other applicable guidance, the standard is more often applied to other types of products and equipment.  

 Although gamma radiation is commonly used to sterilize plastics, not all types of plastics can be treated at a sufficient dose to achieve sterilization without degrading the plastic (20). The assessment of degradation is on-going and should be undertaken throughout the shelf life of the material (the application of ionizing radiation causes the excitation of polymer molecules, where, over time, the adsorbed dose can result in changes to the physical or chemical properties of the polymers). The plastic material to be irradiated is normally referred to as the "product." Most products are placed into an outer packaging in order to protect the irradiated product and to keep the product sterile once sterilized. The product remains sterile provided that the outer packaging remains intact. Occasionally, large or complex products cannot be tested in their entirety and a staged sterilization process is required. In such circumstances, thought should be given to the conditions under which final assembly will take place in order to avoid contamination from the environment or from personnel.  

Given the range of different types of single-use sterilized disposable products being developed, and the range of different packaging configurations, the required gamma radiation dose to achieve sterilization or to protect the product from degradation will vary considerably. There is also considerable variation with types of plastic. For example, a relatively low dose of radiation is required to sterilize polypropylene when compared with polystyrene. Furthermore, the assessment of the dose is more straightforward for small items, such as a plastic container, and more complex for single-use systems. Single-use systems have variables including tubing length, different numbers and types of filters, and differences in the design of containers, bags, and valves, which make the determination of the irradiation dose more complicated. Considering these factors, a common radiation dose used for plastics is in the range 15–25 kGy (21).  

For the process of sterilization, the wrapped product is normally packed into a special container, typically manufactured from aluminum, called a tote. A tote has fixed internal dimensions and is designed to transport product through the radiation process. The weight and dimensions of the tote must be accounted for when establishing the radiation dose. 

The dose determination is the key validation step when using gamma radiation. The dose is the amount of gamma radiation absorbed by an item undergoing sterilization. This is normally set as a range, where a minimum and maximum dose is established. The minimum dose is established as the point where sterilization occurs, and the maximum dose is the point beyond which the product is no longer compatible with the sterilization process. In general, the higher the dose rate, the lower the adverse effects upon polymer products. This is mainly due to the diffusion of oxygen during the irradiation process. 

To validate a load, there are three aspects to consider: establishing the dose range, measuring the effectiveness of the sterilization, and dose mapping.  

 Irradiation validation is designed to set the dose range. The primary focus is to determine if the irradiation process damages the packaging material or the product to be sterilized. Damage is assessed by calculating the maximum dose. This assessment is examined through stability trials, whereby samples are held at under defined storage conditions (i.e., temperature and relative humidity) and examined at periodic intervals for discoloration, brittleness, and other damage.  

Ionizing radiation generates free radicals in plastic polymers leading to degradation from chain scission (i.e., changes in molecular weight) or alterations to cross-linking. Potential radiation effects on some materials include embrittlement (i.e., change to material hardness), discoloration (i.e., often yellowing caused by surface oxidation), unpleasant odor (i.e., from volatile material formed by reactions from within the polymers), or lack of functionality due to a compromised physical trait, such as tensile strength (22). 

 The aim of this sterilization validation is to determine the dose required to achieve a sterility assurance of 106. The probability of sterilization is commonly assessed by bioburden determination (determination of the bacterial and fungal load), as recommended by ISO 11137. This involves the following four steps. 

Determination of the bioburden of the product. This is normally done by taking 10 units per batch from three different batches of product, post-irradiation. In doing so, it is important to be sure that the product that was subjected to gamma radiation was representative of the product normally manufactured. The manufacturer of the product normally carries out the bioburden determination. It is important to ensure that the bioburden recovery method is accurate because insufficient recovery of microorganisms during bioburden tests would result in an underestimation of the true bioburden of the product and lead to an inadequate sterilization dose applied to the product. 

The bioburden of the product is dependent upon several factors. These include the nature and source of the raw material, the components used in manufacturing, the product design and size, the manufacturing process, the manufacturing equipment, and the manufacturing environment (e.g., the type of cleanroom used). For products manufactured using approved suppliers and assembled within cleanrooms certified as ISO 14644 class 7 under localized unidirectional airflow protection, the expected bioburden would be relatively low (e.g., not more than 10 CFU per device). Additional data relating to the risk from the manufacturing environment can be provided through microbiological environmental monitoring. 

There are different methods for bioburden determination. One of the most common methods is the repetitive (exhaustive) recovery method. This method involves washing the sample product repeatedly until it is estimated that no further microorganisms will be recovered. The washing process can include the addition of sterile glass beads or ultrasonication to facilitate microbial recovery. The eluent from the washing should be tested using an appropriate TVC test method (where membrane filtration is the method of choice, followed by the pour plate technique). 

The microbial counts from all washes are compared in order to assess the total bioburden. Such extraction methods require validating. Method validation involves deliberately inoculating a sterile disposable item with a known number of microorganisms and then assessing the number recovered from the washing steps to the theoretical inoculum challenge. A valid method should recover the microbial challenge as per the guidance in 

 <1227> for microbial method validation. Where the challenge cannot be recovered, one factor may be the variation with the process of drying the microbial challenge organism onto the plastic item prior to washing (23). 

Calculation of appropriate dose based on the resistance of an identified microbial population. This is based on the total number of bacteria and fungi isolated and an assessment of the types of species recovered, as characterized using microbiological identification techniques. This is an important distinction as it is not simply the total numbers recovered as some microorganisms have a theoretically greater resistance to gamma radiation processes than others, most notably Streptococcus faecium and Micrococcus radiodurans (24).  

Radiation dose assignment. To select the appropriate radiation dose in relation to the theoretical resistance of the microorganism, ISO 1137 provides a guidance table.  

Validation of calculated dose. As part of the validation, the calculated dose is verified by selecting an appropriate sample size (normally 100 units of the product) to determine if the dose is efficacious. The test is undertaken by placing individual units of product into sterile bottles containing microbiological culture media (e.g., soybean casein digest medium) and incubated for 14 d. This is the "sterility test," although it bears some similarities to the direct inoculation sterility test described in the US or European pharmacopoeias, it is not equivalent (25). Any bottle of media that exhibits microbial growth (turbidity) is indicative of the product not being sterile and that the sterilization cycle is inappropriate for the product.  

As with the bioburden determination method, the test for sterility requires validation. The object of the validation is to show that the product material does not inhibit the growth of microorganisms (a method suitability test). Inhibition of microorganisms would lead to the risk of a false negative result occurring. The method suitability determination involves using the same type and volume of culture media used for the sterility test. The product is inoculated with known numbers of a bacteria culture and a fungal culture. The inoculated product is then incubated. Any product that shows no growth or slowed growth is considered bacteriostatic or fungistatic and the method declared unsuitable (26). Within the medical device industry, an acceptance criterion of a sterility assurance level of 102 has been used (i.e., two units could fail the sterility test and results could be deemed as acceptable). This level of assurance is unacceptable for single-use technologies used in conjunction with aseptically-manufactured products. In such cases, further modification of the method is required, such as increasing the volume of culture media or using culture media with an added neutralizer.  

There are complications with the testing of large products for the bioburden and sterility tests. To carry out tests as commonly practiced, disassembly (i.e., "sectioning") would be required so that the product can be immersed into microbiological culture media (i.e., broth). This is an activity that could result in adventitious contamination due to the need for personnel to manipulate the product. With such devices, an alternative approach is more often adopted that involves sterility of a product's fluid path (i.e., passing a sterile buffer through the product and examining it for microbial growth). The test is assessed by incubating the media in the presence of the product (or product rinse) for 14 days and examining the broth, post-incubation, for microbial growth. 

When assessing the data to determine the final sterilization dose for routine batches of single-use products, it is normal to use the highest dose (Dmax) established in the validation in order to set a level of overkill. This provides additional assurance should the product bioburden shift upwards or should more resistant strains appear.  

A further important consideration is that the bioburden test and sterility test are often carried out by the product manufacturer and gamma irradiation plant, respectively. In order that the data are comparable, it is important that similar test methods, microbiological culture media, and test incubation parameters are employed. If this is not done, then the reporting of a sterility test as zero microorganisms may be incorrect if the sterility test method is not capable of detecting all of the microorganisms detected in the bioburden test. This can arise if different microbiological culture media are employed or if the medium is incubated at a different temperature for a shorter time period. The sterility test result may then be due to the inability of the organism to grow under the test conditions rather than an indication that the organism has been destroyed by the sterilization process.   

Variations to the bioburden method are permitted within the ISO 11137 standard, such as the VDmax25, which permits fewer units of product to be tested and for similar types of product items to be grouped together for the validation (for 25 kGy doses). This variation can only be used where it has been established that the bioburden level is relatively low (at less than 1000 CFU per device). When considering whether different products can be grouped together, account must be made of the plastic materials, construction processes, surface area and handling. 

 The third aspect of the validation is dose mapping. For this, the product, in its final packaging configuration, is profiled in order to identify the high and low zones of absorbed dose in the product load. Dose mapping determines the loading configuration that will be used during routine sterilization. 

 Understanding Gamma Sterilization, by Jerold Martin, Pall Life Sciences 

The object of the validation is to set processing parameters and the product release specification. The validation parameters are established through a performance qualification (dose mappings), which is typically run three times using the maximum packaging size. The main steps for undertaking a performance qualification, for each product, are as follows:  

Establish parameters for routine product sterilization.

The level of radiation is assessed using devices called dosimeters (i.e., a device, instrument, or system that measures the exposure to radiation). It is important to assess the number of dosimeters required to assess the radiation dose. With a standard tote, it is typical to use 15

20 dosimeters. This number is necessary in order to achieve an accurate assessment because the radiation dose applied to the products packaged at the outer edge of the tote is often higher than the dose received by the material in the inner center of the tote, as illustrated in Figure 1. 

The key validation parameters are: product weight and volume, dimensions of packaging components and density, and the configuration of the packaging components. With the dimensions, it is important that the product is evenly distributed because the radiation dose is applied at the same level from both sides. With the issue of load configuration, this point is sometimes overlooked. It is nonetheless important that the way that the components are packaged during validation be replicated for all successive radiation runs. This is important because if the orientation alters, then this can cause changes to the density mix and thus the effectiveness of the irradiation. Once the validation parameters are established, the parameters are used for routine processing and no parameter should be permitted to vary significantly from the established parameter. 

Once established, it is necessary that any future changes in product, its package, or the presentation of product for sterilization are re-assessed for the effect on the appropriateness of the sterilization process. It is prudent to re-assess the validation parameters at least on an annual basis and to assess the bioburden of the product quarterly, in order to determine that the gamma radiation process remains effective (27). This assessment may include fractional studies. Here the product is irradiated and tested at sublethal doses (i.e., at levels of gamma radiation below the minimum established in the validation study) to check for continued dose efficacy. This is sometimes referred to as dose auditing. 

This paper examines the process of gamma irradiation of plastic materials used as part of single-use disposable systems in the pharmaceutical and biotechnology sectors, with a focus on validation requirements.

Application of Sterilization by Gamma Radiation for Single-Use Disposable Technologies in the Biopharmaceutical Sector

Testing for the presence of microbes in an aseptic environment used to be a process that took weeks. Now, methods for rapid microbial testing can be used to detect the presence of mircoorganisms in the environment or in a finished drug product in hours or days. In this technical forum, experts describe different methods of rapid microbial testing and their applications. Contributors to the forum are: Anne Baumstummler, senior scientist; Renaud Chollet, head of biotechnologies and reagents; Hervé Meder, R&D project manager; Céline Rofel, cell culture and molecular biology technician; Adrien Venchiarutti, engineer; and Sébastien Ribault, PhD, director development & bioproduction, all at Millipore SAS; and Ruth Eden, PhD, president of BioLumix. 

Microcolonies Fluorescent Staining Method for Rapid Detection of Microbial Contamination in Mammalian  Cell Culture 

Anne Baumstummler, Renaud Chollet, Hervé Meder, Céline Rofel, Adrien Venchiarutti, and Sébastien Ribault, Millipore SAS

Traditional testing methods for microbial contamination of mammalian cell culture require several days to detect contamination. The presence of microorganisms is typically assessed by membrane filtration and incubation on solid media or inoculation into liquid media. These culture-based methods rely on microorganisms growing and yielding visible colonies or turbidity. Time-to-result is measured in days for standard bioburden testing (1). The microcolonies fluorescent staining method (MFSM) described in this study is a robust, simple, and rapid method enabling the detection of microbial contaminants. It can be applied to cell culture samples with time-to-results that are two to five times shorter than traditional microbiology. The nondestructive feature makes it compatible with downstream identification of contaminants. 

 Many technologies and detection systems such as enzyme-linked immunosorbent assays, impedimetry, bioluminescence, flow cytometry, and polymerase chain reaction arose in the field of rapid detection during the two past decades (2–4). Fluorescent dyes have been routinely used in methods such as the direct epifluorescent filter technique (DEFT) to monitor microbial contamination (5). This technique involves the capture of microorganisms on the surface of polycarbonate membrane filters, staining, and visualization using epifluorescence microscopy. The sensitivity is directly linked to the volume filtered and the number of fields observed. A limitation of this approach is the inability to differentiate fluorescent microorganisms from autofluorescent particles. Automated and semi-automated systems have been developed to increase this discrimination and improve accuracy (6–8).  

An alternative is to combine the principles of epifluorescence microscopy and flow cytometry (9). Microorganisms retained on a polycarbonate filter are fluorescently labeled and automatically counted by a laser-scanning device that makes the differentiation between fluorescent microorganisms and particles (10). Several authors have reported the use of this solid-phase cytometry technology for bioburden testing of mammalian cell culture processes (1). Results are typically obtained within five hours of sample preparation (6). One drawback of such direct detection methods is the inability to discriminate between culturable and nonculturable organisms (11). In addition, many fluorescent-labeling techniques have an impact on the viability of colony formation, thereby making it impossible to identify organisms that are detected. This is a challenge when conducting investigations or evaluating the severity of a contamination.  

MFSM is a method that combines the widely used membrane filtration method with a fluorescence-based staining for the rapid detection of contaminants in filterable samples using microcolony formation. MFSM is based on a nondestructive fluorescent labeling of viable and culturable microorganisms present on cellulose membranes (MilliFlex Quantum, EMD Millipore; see Figure 1). The procedure consists of filtration of the sample, a short incubation on media to yield microcolonies, staining of microorganisms, and counting of fluorescent microcolonies with a light-emitting diode (LED) system. The microorganisms are labeled directly on the filter with a nonfluorescent substrate that is cleaved by intracellular microbial enzymes. Only metabolically active microorganisms with membrane integrity that retain the fluorescent product are stained. Because the staining solution is nondestructive, the method is compatible with standard identification methods. 

The objective of this study was to compare MFSM with traditional epifluorescence microscopy for its ability to rapidly detect microbial contaminants in mammalian cell culture. 

 Both methods were used to detect bacteria in Chinese hamster ovary (CHO) cell cultures. Epifluorescence microscopy was limited by filterability, media interference, and nonrobustness issues, whereas microcolonies fluorescent staining method enabled consistent detection of 

after eight, nine, and 48 hours of incubation. 

With epifluorescence microscopy, nonspiked treated CHO cells yielded a great deal of fluorescent debris and a very high background (see Figure 2A), which interferes with the detection of 

 (see Figure 2B). Because observation was difficult in the presence of cells, 

was spiked in CHO medium. The fluorescent background was removed, but it was still difficult to observe bacteria on the polycarbonate filter. The detection of the microorganisms was only possible when increasing the number of contaminants retained on the filter by tenfold (see Figure 2C). Microorganisms spiked in the mammalian cell culture were easily detected with the MFSM without any background or media interference (see Figure 2, D–F). Background noise was minimized as a result of lysing the CHO cells before the filtration using the mammalian cell lysis solution; this buffer eliminates CHO cells while minimizing the impact on microorganisms. Similar results were obtained for both methods with 

As a result of the nondestructive feature of the MFSM, stained membranes could be reincubated on culture media to yield visible colonies that can be collected for further identification using existing identification methodologies (i.e., biochemical, morphological, nucleic-acid analytics, etc.). The ability to identify the microorganism can accelerate the root cause analysis and implementation of the corrective/preventative action (CAPA) plan.  

  6.  J. Moldenhauer,  (2008) "Overview of rapid microbiological methods" in 

Principles of Bacterial Detection: Biosensors, Recognition Receptors and Microsystems

 M. Zourob, S. Elwary, and A. Turner, Eds. (Springer New York, 2008) pp. 49–79. 

 10. K. Mignon-Godefroy, J.G. Guillet, and C. Butor, 

Growth-Based System for Rapid Microbial Testing

Currently available growth-based rapid microbiological methods provide either a quantitative cell count, estimation of viable cell concentration, information regarding the presence of a specific microorganism, or a microbial identification. However, the ability of these rapid microbiological methods technologies are limited in scope in that they cannot be used to perform all the required assays using a single technology platform. A new automated growth-based system simultaneously detects microbial growth, provides an estimation of viable cell counts, and identifies the presence of specified micro-organisms. 

 The system is based upon detection of optical variations due to microbial metabolism in liquid medium within a two-zone ready-to-use test vial. An optical sensor monitors optical changes within the vial's reading zone, which is physically separated from the incubation zone. This two-zone approach prevents masking of the optical pathway by product or microbial turbidity and, therefore, eliminates product interference. Separate test vials are used to automatically detect the presence of viable microorganisms and to estimate the concentration of viable counts by monitoring changes in CO

 production during cellular growth. The CO

  sensor is composted of a matrix of a polymeric material that is transparent to light and contains an indicator agent sensitive to CO

  gas generated by micororganisms, changing the indicator color from blue-green to yellow. Alternatively, the hydolysis of fluorogenic synthetic substrates by bacterial enzymes can cause an increase in fluorescence. Also, pH changes can be monitored calorimetrically. Each of these applications can be simultaneously performed using the same instrumentation and at the same time.  

The sensitivity of the system is a single viable cell per sample vial; when a single cell replicates to a specific detection threshold level, a positive response is recorded. The threshold level is ~100,000 cells/mL for bacteria and ~10,000 cells/mL for yeast and molds. Additionally, the system yields significantly faster results than the plate count method; one bacterial cell is usually detected within 8–18 hours, a single yeast cell is detected in 20–30 hours, and mold cells require 35–48 hours.   

The system creates dynamic patterns as the microorganisms grow in the medium. As shown in Figure 1, the green curve shows a pattern where no growth had occurred. The curve is flat without any significant increase in the signal and no detection time (DT) is observed. The blue curve shows the pattern generated when the microorganism grows in the vial (a DT of 11 hours is observed). 

In highly contaminated samples, bacteria are typically detected in 8–12 hours, yeast in 16–24 hours, and mold in 24–35 hours, providing timely warning of contamination. 

 Each BioLumix instrument has a capacity of 32 sample locations with a single incubating temperature. Multiple instruments (up to 32 instruments) can be attached to a single computer with Windows-based program that controls the operation of the instrument(s) and is barcode capable. The software is validated to meet 21 

 Part 11 requirements, provides an audit trail, operator identification (log in and log out), trend analysis, and provides various data reports. Detection events are automatically displayed. 

A critical element of the technology is the two-zone detection vial with an upper incubation zone where the sample is added and a lower reading zone that remains optically clear and free of turbidity from microorganisms and sample components. This two-zone vial design eliminates interference of the optical pathway during color and fluorescence monitoring by the sample and microbial growth.   

The data generated for the comparison of the growth-based rapid method to 

Microbial Examination of Nonsterile Products: Microbial Enumeration Tests

, is summarized in Table I. The table shows the specification ranges tested, the number of samples, and the percent agreement between the new method and the 

 plate count method using soybean casein digest agar. 

(i) Naturally contaminated samples: Two hundred and one nonsterile over-the-counter medicine samples such as vitamins, antacids, suppositories, laxatives, ibuprofin, and aspirin were tested. All noninoculated samples were below the detection level by both methods. There was 100% agreement between the two methods in classifying samples as above or below the specification levels. This indicates the ability of the BioLumix system to yield equivalent results to the plate count method when detecting growth over the range of 10 cfu/g to 1000 cfu/g.   

(ii) Inoculated products: Fifty-nine products were inoculated with all the organisms cited in 

 <61>. Ten inoculated samples had counts below the specification level and all were correctly classified as the challenge organism by the BioLumix system. There was 100% agreement between the two methods in determining whether samples were above or below the specification level.  

As shown in Table I, similar data were obtained for yeast and mold as well as Gram negative bile-tolerant bacteria. The data indicate the ability of the system to yield equivalent results to the plate-count method over the range of 10–1000 cfu/g for both of these assays. 

The method has good specificity in detecting target organisms and excluding nontarget flora, and the detection limit for the system equals or is slightly better that the limit for the plate count method. High precision or repeatability was obtained for all three assays tested. It can be used to detect the presence or absence of organisms, total aerobic count, the presence of yeast and mold, enterobacterial count, and absence of objectionable organisms in 10 grams of product. By encompassing both 

types of testing, the system offers a complete screening solution, making the existing microbial testing simpler, faster, and automated.  

In this technical forum, experts describe different methods of rapid microbial testing and their applications.

Rapid Microbial Testing

PharmTech: What is the difference between parametric release and real-time release testing? 

 Compliance with release specifications can be demonstrated by performing a complete set of tests according to approved specifications before the final product is released onto the market. Under certain conditions, however, it is possible to use an alternative strategy called parametric release. Rather than conducting an approved sterility test, parametric testing involves utilizing information collected during the manufacturing process to demonstrate that the product was made according to defined GMP and sterilization procedures (1). 

Parametric release can only be applied to products terminally sterilized in their final containers by steam, dry heat or ionizing radiation, but recent guidelines adopted in the International Conference on Harmonization (ICH) context (ICH Q8, Q9, and Q10 Guidelines) now enable a similar release strategy to be adopted for other tests as well (2). This approach is called real-time release (RTR) testing, and can be applied to both new and established products. As with parametric release, RTR testing provides assurance of final product quality based on information collected during the manufacturing process. The basis for establishing RTR testing mechanisms involves combining quality riskmanagement principles, enhanced product knowledge and process understanding, as well as applying an adequate pharmaceutical quality system. It is important to note that the other products mentioned may be chemical products such as excipients, active substances, pharmaceutical intermediates and finished products. 

To demonstrate, a combination of in-process tablet weight, blend content uniformity measurement, drug substance purity and particle size could serve as a control strategy for ensuring the drug content of a highdose tablet, if the relationships has been demonstrated. In this example, the above-mentioned factors would serve as the RTR tests. Based on the outcome of these tests, as well as other required tests in the product specification, and GMP compliance, the production batches are released by the qualified person.  

PharmTech: What preconditions have to be given for parametric  release of sterile drug products?

Parametric release is defined by the EMA as a sterility assurance release program where a demonstrated control of the sterilization process enables a company to use defined critical process controls in lieu of the sterility test to fulfil the intent of 21 

Compared with performing sterility tests of units drawn from a finished batch, meeting parametricrelease requirements can provide greater assurance of a product's sterility. Traditional sterility testing methods are limited in their ability to detect contamination because the small number of samples required restricts the ability to capture microorganisms dispersed in large volumes. Additionally, the prescribed culture media has only a limited ability to stimulate the growth of all potential microorganisms. The sterility tests described in the 

, however, provide an optimized approach by taking into account different culture media, and their respective incubation temperatures and incubation times. Even so, sterility tests typically only detect major errors in the manufacturing process that stem from the contamination of a large number of product units. Data derived from in-process controls of a validated terminal sterilisation process, on the other hand, can provide more accurate information regarding product sterility because the probability of product bioburden surviving the sterilization process in any single unit of a product can be calculated to less than one in a million. 

At the beginning, the accurate monitoring of relevant sterilisation cycle parameters, such as temperature, pressure, time and the fθ value, were sufficient for a parametric release. However, further requirements soon became mandatory. Today, the entire sterilization process must be described, and process validation reports have to comprise heat distribution and heat penetration studies for at least three runs for each load pattern used, according to good validation practices. Microbiological qualification must also show sufficient efficacy at the minimum level of the cycle, including information on the biological indicators used (type, D-value, Z-value and stability) and bioburden characteristics (number, type and resistance), as applicable. 

The effort involved in the preparation phase of parametric release is high and a large amount of documentation is necessary, but in most cases, the same effort is also required when following the usual application processes for sterilized drugs. Sterility tests require large additional investments, but also offer additional protection in case of legal claims. The biggest advantage of using parametric release, besides quality assurance, is that it can shorten processing times by a few weeks. Depending on production costs, batch sizes and the amount of batches released, this can lead to significant financial savings. However, manufacturers also need to bear in mind that not all countries accept parametric release.  

PharmTech: Will real-time release testing replace the final release procedure? 

The main benefit of RTR testing when combined with quality by design and process analytical technology (PAT) tools is increased quality assurance for patients. In addition, such a strategy can lower manufacturing costs through improved yields, less waste, faster cycle times, and fewer deviations and rejects. 

Process understanding and the definition of a robust control strategy are key aspects of RTR. The interaction of material parameters and attributes, as well as the dynamics of processes, must be taken into account, and are one important part of submission dossiers for marketing authorisations. As such, when implementing RTR systems it is essential to use PAT to efficiently monitor and record all quality-relevant parameters. In this area, there have been a number of improvements, including advances in NIR- and Raman microscopy, fluorescence microscopy, image analysis and laser-based methods. Using chemometics and multivariate analysis, it is now also possible to monitor production processes in real-time. However, a move towards RTR is not only about using advanced analytical and physical measurement techniques, but also involves performing statistical calculations on the resulting data masses. Importantly, regulatory authorities worldwide have to accept the move from final product testing to RTR applications. RTR has to be accepted by all regulatory authorities because the current varied acceptance can be a major issue. The equivalence between RTR-testing and pharmacopoeial testing must also be explained.  

If one wishes to implement new tests and procedures, then it is important to adapt the whole quality assurance system. This approach is mandatory for compliance. The new quality system must also be aligned with ICH Q10. 

These steps constitute the transition from rigid to risk-based cGMP, and a mindset change from reactive to proactive quality thinking, which could be quite a challenge. In particular, there will be a huge impact on qualified persons or other persons responsible for product release. Specifications will need to be elaborated carefully, and the definition of deviations that require investigation and the process to declare analytical results as outliers or invalid data must be described in SOPs. 

RTR will be implemented over time as industry understanding increases and regulatory expectations are clarified. There is a long road ahead, but as the saying goes: a journey of a thousand miles begins with a single step. 

is head of quality systems excellence at Boehringer Ingelheim GmbH (Ingelheim), 

 1. EMA, Note for Guidance on Parametric Release (March 2001). 

 3. 21 CFR Part 11 Title 21 (April 2010).  

Parametric release and real-time testing use manufacturing data to ensure that products are made according to defined standards. PharmTech talks to Boehringer Ingelheim's Heribert Hausler about these issues.

Parametric Release and Real-Time Release Testing

For decades, two different approaches have been used to manufacture sterile products. The most common approach, aseptic processing, relies upon separating potential microbial contaminants from the product through a combination of filtration of the product stream, individual sterilization of product-contact packaging components, and environmental controls. Terminal sterilization relies on a more limited contamination control of the product, components, equipment, and environment followed by a lethal sterilization process applied to the fully assembled dosage form. From a macro perspective, these two approaches have been treated as equivalent means to the same end, but they have never been considered equal in terms of process capability. In the jargon of sterile-product manufacturing, they are not considered fully equivalent in terms of sterility assurance. 

The world's regulatory authorities have long expressed a preference for terminal sterilization, but have rather inexplicably set up expectations that provide little real benefit to firms that implement terminal sterilization. The foremost shortcoming is requiring firms to make a regulatory filing to delete the compendial sterility test as a requirement for product release, a practice known as parametric release. Regulatory pressure exists to manufacture products for terminal sterilization in ways increasingly similar to aseptic processing. This change involves added facility controls, stricter gowning requirements, and substantial environmental monitoring that further deteriorate any financial benefit that would accrue to firms using terminal sterilization. In Europe and to a lesser extent in the United States, another blow against terminal sterilization has been the increasingly commonly enforced idea that terminal sterilization should require lethalities in the range of F

 refers to lethality in moist-heat sterilization; by convention, F

 is equal to 1 min of exposure to moist heat at 121.1 °C.) 

In Japan, a different and far more scientifically logical approach has evolved and is undergoing further refinement. The idea is that with reasonable control of presterilization bioburden, lower levels of lethality can afford high levels of patient safety. Specifically, for reasonably heat-stabile products F

 values of 2 min or less can provide levels of end-user safety substantially higher than those attainable in aseptic processing. The logic of this approach is irrefutable as can be shown by simple microbiology. 

Although spore-bearing gram-positive rods with substantial heat resistance are used to develop and validate sterilization cycles, the most common human and animal pathogens are bacteria, mold, and virus with substantially lower heat resistance than these spore-bearing organisms. Protein, a key structural and functional component of microorganisms, is typically denatured in vegetative mold, bacteria, and virus at temperatures only slightly greater than 56 °C. Unsurprisingly, these organisms die quite readily at temperatures in the range of 65–75 °C. This difference means that substantial increases in user safety could accrue at temperatures that are substantially lower than those at which spore killing begins and at which F

 begins to accumulate, which is in the range of 100–105 °C. Thus, the vast majority of medically significant organisms would be dead well before a postassembly heat process reached F

 = 0.1 min. The improved patient safety that this approach could add to moderately heat-stable products is so obvious that one wonders why it is so steadfastly ignored. In fact, a process need not get to 100 °C to add substantial safety, five minutes or less at 70 °C or so would be appropriate. 

One used to applying the common pharmaceutical approach to sterilization might ask whether a process that did not kill spores had value? Aseptic processing is a completely nonlethal process and yet is widely accepted. Of course, a moist-heat sterilization of F

 = 2 min would effeciently kill many spores of many spore-bearing species, including those known to produce human or animal disease. (The recommended means to deliver this process is at temperature less than 121 °C, whereby more uniform conditions can be delivered through the use of a much shorter time at an elevated temperature). Most spores isolated from industrial environments have D

is the time required to reduce a microbial population by 90% when exposed to moist heat at 121 °C.) Given the conservative assumption that the presterilization bioburden following an aseptic processing would be no more than one organism per container and the organisms present had a D

 value of 0.2 min, a process yielding an F

 = 2 min would produce a probability of nonsterility of 10

. In medical terms, this calculation dramatically understates the actual safety because the majority of the bioburden (especially those considered human pathogens) would be non-spore forming and would therefore be dead while the process was still in heat-up mode. 

Dr. Tsuguo Sasaki, GMP Expert, Office of Compliance and Standards of the Japanese Pharmaceuticals and Medical Devices Agency (PMDA) advocates that F

 ≥ 2 min would be a reasonable expectation for parametric release. The authors strongly agree and would favor a harmonized regulatory approach in which any firm with acceptable GMP compliance that submitted an application for a product sterilized at a process lethality of F

 ≥ 2 min would be granted parametric release for its product without any added expectations. This proposal is radical only because existing regulatory policy fails to properly consider true medical risk. 

Parametric release simply means releasing product without having subjected it to a compendial sterility test. The real question is what value the performance of a sterility test on a process offers. Given the low sensitivity and statistical sampling limitations of the sterility test, it has limited value and really should be called a test for gross product contamination. It is rarely failed when applied to aseptically produced products and simply has no value when used to test products sterilized in their containers. 

Lower temperature heat treatments would provide significant safety benefit to products that are filled in human-scale cleanrooms. The benefit of these treatments would diminish in instances where advanced aseptic processing technologies, such as isolators, closed restricted access barrier systems, or some blow, fill, and seal processes are used. The authors postulate that comparable approaches could be used with radiation sterilization where low-dose exposures would provide similar advantages to patient safety. ISO 11137-2 provides for developing relatively low does processes based upon bioburden number and resistance (1). 

Recommendations and experience from Japan

PMDA's Dr. Sasaki shared with the authors some basic concepts from Japan's 

Guideline on the Manufacture of Sterile Drugs by Terminal Sterilization

, which is expected to be issued in 2012. This document will consist of a narrative section and annexes. The following information reflects general recommendations for low F

 terminal sterilization and will be presented in one of the guideline's annexes:  

Cleanliness of filling area is Grade A, and surrounding areas are Grade B. (Grade C may also be used but personnel microbial monitoring would be expected to meet Grade B levels.)

Average presterilization bioburden should be less than 1 colony-forming unit (cfu)/product unit.

Bioburden should be evaluated for heat resistance.

Grade A is used for filling, but media fill testing is not required.

Low-dose terminal sterilizational is in widespread use in Japan. Dr. Sasaki provided some data regarding its application in Japan during 2005–2010 from 13 different manufacturers (see Table 1). Nearly 80% of the roughly 2.8 trillion units manufactured by terminal sterilization were subjected to F

 of two minutes or less. Quite clearly, these low lethality sterilization processes would not be allowed were they not safe and efficacious. 

 Table I: Application of low-dose sterilization from 13 manufacturers in Japan from 2005â2010.  

The re-examination of the regulatory philosophies that underpin sterile-product manufacturing is long overdue. There is a failure to recognize that postfill heat and radiation treatments could provide additive safety benefits and that these adjuncts to aseptic processing may be applicable to a significant range of products. A process need only be as lethal as necessary to destroy the bioburden present. The current gulf between aseptic processing and terminal sterilization can be bridged as the information from Japan clearly illustrates. At the same time, these data indicate that true terminal sterilization can be applied far more frequently than it is currently by the industry if one considers that low lethality processes can yield safety levels that are so good that further improvements would provide only theoretical medical benefit.  

What has to happen other than this re-examination of the fundamental regulatory philosophies for sterile-product manufacturing? The authors suggest the following:  

 The realization that most medically significant organisms die at temperatures and conditions far lower than where modern industry considers that sterilization begins.

 principal has tremendous value and relevance and that there is nothing magical about 121.1 °C as a minimum target temperature for sterilization. 

. Increased awareness that parametric release should require evidence only that a process does not benefit from the application of the compendial sterility test as a quality-release requirement. 

. Awareness that Item 3 requires disavowal of the notion that there is a test to prove sterility or that such a test could be devised. No microbial test, be it growth-based or rapid, has a limit of detection of zero. It is impossible to prove a negative absolute.

. Understanding and accepting the clear scientific principal that neither safety nor sterility hinge upon the ability to kill 10

spores per article. Organisms isolated from extreme environments, such as hot springs and other geothermal features, are not and cannot be medically significant. 

. We can learn from the extensive experience of Japan with respect to lower lethality terminal sterilization.

The authors urge broad international discussions on sterile- product manufacturing and strongly suggest that these discussions be undertaken with the understanding that patient safety must be evaluated from a medical perspective only and not from the viewpoint that begins and ends with current regulatory paradigms. Many modern-day sterile-product manufacturing paradigms are mistaken and unduly inflexible. The result is a stifling of innovation and an overreaction to misperceived risk while thwarting simple steps that would add safety and reduce cost. 

The authors thank PMDA's Dr. Sasaki for the information he provided for this article and for the many discussions that have been instrumental in exploring the options examined in this article. 

is president of Akers Kennedy and Associates, PO Box 22562, Kansas City, MO, 64113, 

is president of Agalloco & Associates. He is also a member of 

*To whom all correspondence should be addressed. 

Sterilization of Health Care Products. Radiation 

The authors assert that the current gulf between aseptic processing and terminal sterilization can be bridged by re-examining fundamental regulatory philosophies for sterile-product manufacturing.

A More Rational Approach for Sterile Product Manufacturing

Aseptic filling remains a risky process with multiple contaminations occurring each year that have serious consequences. Analysis of an outbreak database shows that, among 1537 patients contaminated by parenteral product from 1990–2005, the mortality rate was 15%. While the majority was due to product preparation at the hospital pharmacy or the practices of the hospital staff, 20% of these contaminations were due to the pharmaceutical manufacturing process (1).  

This high contamination risk leads authorities to regularly reevaluate their requirements, thus making aseptic filling one of the most complex processes in the pharmaceutical industry. A continuous improvement approach, however, has led to major innovations, and the rate of contamination has been significantly reduced during the past 50 years. The innovations listed below involve both the container and filling technologies: 

 The vial emerged as the standard primary packaging and replaced the old-fashioned ampoule, which had higher risk of breakage, contamination through small cracks, and glass particle generation when the ampoule was opened.  

Better practices have been developed for aseptic filling, such as personnel gowning, equipment/process design, and environment monitoring. 

Physical barriers have been created between the operator and the filling area. Initially the barrier was made from simple walls but now includes the isolator concept. Isolators prevent direct contact between the operator and the filling process, including the bulk products, containers, and product contact parts (2).  

Process analytical technology (PAT) was developed to perform on-line checks on product quality. Checks include the classical weight check as well as newer checks such as particle inspection and leak detection.

These innovations improve product quality but also add complexity to the aseptic-filling process. As a result, aseptic filling is expensive, demands complex quality control, and has many potential opportunities for mistakes.  Closed-vial technology, however, can improve product quality and simplify the aseptic-filling process (3, 4).  

Figure 1 shows an overview of the production and filling of closed vials, which occurs at three separate facilities.   The vial is molded and assembled in an ISO5 cleanroom and rings are added.  At a separate facility, the vial is sterilized by gamma irradiation, which leads to a clean, sterile, ready-to-fill vial.   The vial is delivered to the pharmaceutical filling site for filling.  In this process, a needle punctures the stopper and dispenses the liquid. The puncture trace is  then resealed with a laser to restore the closure integrity. Finally, the vial is capped with a snap-fit, polyethylene cap. 

 Figure 1: Production cycle of a closed vial from molding the container to filling and capping.  

Closed vials can offer three main advantages compared with traditional glass vials. 

 In glass vial technology, the vial stays open for more than 30 minutes between exiting the depyrogenation tunnel and stoppering. Stoppers may remain in a stopper bowl for several hours, in which direct contact with surfaces increases the risk of transferring a contaminant to the vial.  A closed vial, however, remains permanently closed except during needle penetration, thereby reducing the risk of contaminant entering the vial by two logs (5). 

 The closed vial is delivered clean and sterile, allowing the pharmaceutical manufacturer to eliminate container-component preparation, including water for injection (WFI) washing, steam sterilization, and hot-air depyrogenation. High speed stoppering and aluminum cap crimping are also eliminated. The break-resistant polymer material reduces vial breakage inside the filling area and during the supply chain. 

Easier handling for healthcare professionals.

 The closed vial's cap can be easily opened by breaking small polyethylene bridges. Piercing is facilitated by a large piercing area. Liquid collection is complete due to the absence of recess areas in the stopper design. Finally, the vial does not break if dropped. A market study performed for Aseptic Technologies in 2007 found that among 246 professionals (i.e., medical doctors, nurses and hospital pharmacists), 87% preferred the closed vial (Crystal Closed Vial, Aseptic Technologies) and 7% preferred the glass vial. The most often cited reason for preferring the closed vial, as shown in Figure 2, was that it is easy to handle. 

Figure 2: Results of market survey of healthcare practitioners showing the reason for their preference of closed over glass vials.  

The following sections describe a typical closed-vial container and the manufacturing and filling process. These sections also explain how the container design and the manufacturing process provide a solution for challenges in aseptic filling. 

In this closed-vial system, cyclo-olefin copolymer (COC) (Topas, Topas Advanced Polymer) was selected for the vial body because it does not create high particle levels during molding.  Low particle generation is a requirement for avoiding WFI washing after manufacturing in an ISO5 clean room. COC is already used in some injectable products (Metalyse, Boehringer-Ingelheim), and is widely used in blister packaging. COC is a clear, transparent polymer that allows good light transmission and has a high barrier to water vapor.  In addition, it can be gamma-irradiated without degradation or a visible change of color at standard irradiation doses. COC is shock resistant, which reduces the risk of loss during production and transportation. 

Polymer molding has greater design flexibility compared to glass forming.  Several features are shown in Figure 3. In particular, the tightness of the vial is ensured under all conditions, even under the low temperatures of liquid nitrogen. 

Figure 3: Detail of closed vial container design showing features not available in glass vials. 

The stopper should reseal when heated by the laser to ensure reclosing of the puncture trace.  The stopper must be able to absorb the laser energy with a good profile of heat distribution. Second, the stopper should be highly flexible and easy to pierce with a large needle without generating particles of significant size or amount and without material loss. Third, to ensure optimal resealing process after liquid fill and after lyophilization, the stopper should have good elastic memory. It is crucial to have both sides of the piercing trace in tight contact to ensure optimal laser resealing. Finally, the stopper material used should not release deleterious leachables.  A thermoplastic elastomer (TPE) has these features, and the polymer can be engineered using a color pigment to ensure optimal absorption of laser energy.  

 The vial head is equipped with a top ring to secure the assembly of the vial body and the stopper, as shown in Figure 3. In this design, the vial head has also been equipped with a snap-fit, high density polyethylene (HDPE) cap.  This design eliminates the complex and particle-generating crimping process necessary with an aluminum cap. A small rib on the internal surface of the cap adds closure integrity by isolating the central part of the stopper from the environment until use by the doctor.  

Closed-vial manufacturing process description

The major innovation of the closed-vial technology is the production of the vial components in an ISO5 clean room. As a result, the components are ready-to-use and do not require the complex cleaning process that is mandatory for glass vials and rubber stoppers.  

 To ensure the cleanliness of the vial components, various conditions have been imposed on the process. First, the molds should not contain lubricating additives that are sometimes used to ease removal of the part from the mold.  Second, once the room is qualified for operation, the operators cannot enter. Vial conveying must be fully automatic and should not create particles above the specified level.  Vial transportation is performed by robots as shown in the first step of Figure 1. Such robots are already widely used in the ISO4 or ISO3 cleanrooms in the electronic industry as well as in the pharmaceutical industry for applications such as syringe filling.  

Because the robots can achieve high precision, the stopper is designed to come straight to the vial body, which avoids the presence of a recess area when the vial is upside-down for liquid collection. Therefore, all the liquid will reach the bottom of the vial and be collected (see Figure 3). As a result, vial overfill can be reduced, thus leading to significant savings of API. 

After assembly of the two components, top and bottom rings are added. Each step is checked by visual sensor control or mechanical challenge before moving to the next operation.  This complete PAT ensures that the vial is fully and properly assembled.  

 Because the TPE used for the stopper is sensitive to heat, the only classical sterilization procedure suitable for the closed vial is irradiation. Gamma irradiation is preferred to beta irradiation because it is available worldwide and can process a complete pallet at once.  

Closed vials are provided ready-to-fill.  The five most frequently used methods for loading are: 

Wrapped vials are loaded into the filling area before sanitizing so the external part of the bag is sanitized along with the equipment. This method is used  for very small batches (i.e., maximum of a few hundred vials).  

Beta-bags are connected to rapid transfer ports. This method is used  for small batches (i.e., a few thousand vials) with robot filling lines using closed vials in racks. 

Vials enter through vaporized hydrogen peroxide airlock with sanitization of the last bag. This method has a limited capacity, due to either the small size of the airlock or the long cycle time, and is therefore efficient for low-capacity filling equipment. 

Vials entry through airlock cascades from an ISO8 to ISO5 environment, using robots to perform automatic debagging and box opening. This is used for mid to large scale batches (e.g., 25–200 vials/min.). 

Vials are removed from bags and boxes, followed by e-beam sterilization of the stopper top surface.  This method is used for very large batches (i.e., up to  600 vials/min.).

Using ready-to-fill vials eliminates component preparation and thus has a huge impact on the entire facility.  Equipment for vial washing, a hot-air tunnel, and equipment for stopper washing/sterilization is not needed, and clean room space is reduced.  WFI for formulation and equipment cleaning can be sourced from a much smaller WFI loop, or containers may be purchased from external sources.  

Another change to the filling process is that a needle must pierce the vial stopper before filling, and the hole must be reclosed after filling. The vial must be held in a fixed position during piercing, filling, and removal of the needle. The vial must also be held in position under the laser head to be resealed, and the laser must ensure complete coverage of the piercing trace, so the laser has a uniform energy beam on a 6 mm diameter surface. After resealing, a snap-fit cap is pressed in place. 

The closed vial acts as a mini-isolator because exposed surfaces are limited to the stopper top surface and the needle. In contrast, in a traditional glass vial, the inside of the vial, the inside surface of the rubber stopper, and the needle are exposed until the vial is stoppered.  The closed vial filling system (CVFS) offers a new barrier or containment concept, in which only closed containers are handled (6, 7). The CVFS   is suitable for installation in an ISO8 cleanroom, as illustrated in Figure 4, in which a robotic filling line is surrounded by a CVFS. 

Figure 4: A closed-vial filling system surrounding a robotic filling line.  

The advantage of the CVFS over the traditional isolator is its simplicity, in that it can be sanitized with classical sporicidal agents and does not require vapor hydrogen peroxide sanitization. The CVFS uses unidirectional, HEPA-filtered laminar airflow that exits through the bottom of the system, which helps maintain laminar flow and prevent turbulence.  Isolators are still mandatory when the safety of the operator must be ensured (i.e., with highly potent drugs such as cytotoxics). Such isolators must be installed in an ISO9 cleanroom.  

The advantage of the CVFS versus the Restricted Access Barrier System (RABS) is that in the CVFS,  operator access is only possible via gloves, and the barrier environment is never compromised by door opening.  Material entry is limited to secured processes such as rapid transfer ports, airlocks, and e-beam irradiation units. With these limitations, an ISO8 clean room environment for the surroundings is sufficient to ensure the ISO5 quality inside the barrier. 

Closed-vial technology is an alternative to traditional glass vial filling that reduces the risk of contamination for the patient, simplifies the filling process, and provides easier handling for healthcare providers.

Pharmaceutical Technology-05-01-2012

Pharmaceutical Technology, Bioprocessing Supplement 2012 (PDF)

Application of Sterilization by Gamma Radiation for Single-Use Disposable Technologies in the Biopharmaceutical Sector

Rapid Microbial Testing

Parametric Release and Real-Time Release Testing

A More Rational Approach for Sterile Product Manufacturing

Using Closed-Vial Technology in Aseptic Filling

Biological Indicator Growout Times: The Impact on Establishing a Reduced Incubation Time Protocol

Bioprocessing Trends: Annual Survey Results