Releasing capacity of pre-sterile cotton swabs for discharging sampled microorganisms

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1 European Journal of Parenteral & Pharmaceutical Sciences 2016; 21(4): Pharmaceutical and Healthcare Sciences Society Releasing capacity of pre-sterile cotton swabs for discharging sampled microorganisms Ravikrishna Satyada 1 and Tim Sandle 2 * 1 Microbiology Manager, Gland Pharma Limited, Hyderabad, India 2 Pharmaceutical Microbiology Interest Group, Stanstead Abbots, Hertfordshire, UK Surface monitoring by using swabs forms a regular part of environmental monitoring of cleanrooms. There are different factors that affect swab recovery, from tip type to enumeration method. One factor with swabs where the microorganisms are detached from the swab tip and which are then membrane filtered is the period of vortex mixing. This paper discusses microbial surface sampling, and the factors that affect swab recovery, and presents some experimental data where vortex times are considered for a range of microorganisms. The study outcome indicates that 15 seconds of vortex mixing is sufficient to obtain microbial recoveries from the swab tip above 50%. Key words: Environmental monitoring, swabs, microbiology, cleanrooms, method validation. Introduction Cleanrooms and controlled environments provide the surrounding structure within which pharmaceutical products are manufactured. The most important factor achieving environmental control is through good cleanroom design. This includes high-efficiency particulate air filtered air, appropriate air change rates, good air distribution, and the maintenance of pressure differentials. Once control has been established, the assessment of cleanrooms during pharmaceutical manufacturing provides a useful detection method, given the limitations of environmental monitoring methods. Established techniques for environmental monitoring include particle counting and culture-dependent methods, such as active air sampling, settle plates, and surface monitoring through the use of contact plates and swabs 1. To these a new generation of environmental monitoring methods can be added, such as real-time spectrophotometric counters that differentiate between inert and biologic particles 2. Moreover, some alternative environmental monitoring methods are available, such as ATP-based swabs, although the sensitivity is often not suitable for the pharmaceutical sector 3. The objective of viable monitoring is to detect levels of bacteria and fungi present in defined locations within the cleanroom during a particular stage of processing. Excursions, either as counts above pre-defined action levels or a frequency of excursions, can be trended, investigated and risk assessed 4. Contaminated surfaces present a risk should contamination transfer occur, such as an operator placing a hand onto a *Corresponding author: Tim Sandle, Pharmaceutical Microbiology Interest Group, Stanstead Abbots, Hetfordshire, UK; timsandle@btinternet.com contaminated surface and then proceeding to undertake an activity within a clean area; or where a utensil makes contact. Thus environmental monitoring of surfaces is important to assess the numbers and types of microorganisms present 5. Surface monitoring can be differentiated between floor surfaces and surface at working height. Surfaces at working height generally are considered to be more critical as they relate to a preparation step or product contact area. Samples from floors are less critical since no product touches the floor, and the monitoring of these locations acts as a check of cleaning and sanitisation practices. Action levels are set based on regulatory guidance (based on equivalency to the limit for contact plates), with alert levels based on historical data that are reviewed periodically. To monitor surfaces, proven methods are required, such as easy-to-use contact plates, contact slides or swabs. These are, as base concepts, longstanding environmental monitoring methods 6. The contact plate method is recommended when quantitative data are sought from flat, impervious surfaces. Contact plates are filled so that the media forms a dome. The nutrient medium used in the contact plate may also contain a neutralising agent to act against any disinfectant residues that may be present. With the plate, the surface of the media is pressed against the surface being tested for a period of not less than 5 seconds (sometimes longer times are necessary). The resulting sampled area for a 50 mm plate is approximately 25 cm 2. The plates are incubated for the required amount of time, and colony forming units (CFU), if present, are then counted. Swabbing is employed for equipment and irregular surfaces where contact plates are not suitable. Swabs vary in the materials used for the applicator stick (either wood or, more often, plastic) and the tip. Types of swabs that may be used for the swabbing technique include cotton, synthetics and calcium alginate materials with the appropriate diluent. 121

2 122 RAVIKRISHNA SATYADA, TIM SANDLE The swab method involves obtaining the sample by rubbing and rotating a sterile swab, moistened with a nonnutritive or resuscitation medium in several directions over a standardised sample area. Often 5 cm x 5 cm is prescribed to mimic the area sampled by a standard size contact plate, and to linearise data with recommended EU good manufacturing practice (GMP) action limits for contact plates. The swab is then placed back in to the tube containing the medium and agitated or vortexed to transfer the microorganisms present on the swab into the solution. The collection medium may be tested by a most probable number method, membrane filtration method, or direct plating method. An alternative swab, suitable for EU GMP Grade A/International Organization for Standardization (ISO) 14644:2015 Class 5 environments where any contamination presents a risk to an aseptically filled product, are those where the tip is deposited into liquid culture media 7. Swab recoveries are generally considered to be poor, although recoveries can be improved based on the design of the swab and through standardising the swabbing technique (pressure applied, number of strokes, swab rotation and area sampled are examples of variables that can be controlled) 8. With design, there is a difference in recovery performance between the tip material (such as whether it is made from Nylon TM, Raylon TM, cotton, or an alternative material like a cellulose viscose sponge material) 9 ; and, significantly, between plain swabs and flocked swabs. With plain swabs, typically the recovery is only 5 15% from the swab tip to an agar plate 10. However, with flocked (woven) swab tips, recoveries of between 40 and 60% from surfaces are possible 11,12. The tip of the applicator of flocked swabs is coated with short fibres arranged in a perpendicular fashion. The perpendicular arrangement (flocking) is formed when the fibres are sprayed onto the tip of the swab while the tip is held in an electrostatic field. Recovery is superior because capillary action between the fibre strands facilitates strong hydraulic uptake of a liquid sample, including any microorganisms present 13. The microbial recovery from a contact plate is superior to that of the swab 14,15, and, therefore, the contact plate should preferably be used (with swabs used where contact plates cannot be used, such as on narrow or irregular surfaces) 16. A second advantage with the use of the contact plate is that it involves one less step at which adventitious contamination can occur (the contact plate is used and the lid replaced, whereas the conventional swab, depending on its design, either requires plating out or placing into liquid which then requires filtering). However, swabs, based on cleanroom and equipment design, play an important part of the cleanroom environmental monitoring programme. With standard swabs, in order to demonstrate that the recovery from the swabs is valid, users should perform a recovery study along with proving the vortexing period is suitable. The vortex mixing is a key aspect in releasing the sampled microorganisms, and thus it directly impinges on recovery. This paper outlines a study designed to establish the releasing capacity of sterile cotton swabs to discharge the sampled microorganisms. This was based on a variation of the vortex times to see whether 15, 30, 45 or 60 seconds affected the recovery. Materials and methods The swabs used in the study were manufactured by Hi- Media (Mumbai, India). The specification was: sterile cotton flocked swab with a polypropylene stick, held in a polypropylene tube (150 mm x 12 mm). For the study, the following materials were used. Soya bean casein digest agar (equivalent to tryptone soya agar) Sterile saline (0.9% NaCl) Sterile forceps Sterile membrane filtration units Sterile membranes (0.45 µm, 47 mm diameter) Vortex mixer Manifold Swabs Micropipette and tips Stop watch Microbial cultures, sourced from the American Type Culture Collection (ATCC), used were as followed. Staphylococcus aureus ATCC 6538 Pseudomonas aeruginosa ATCC 9027 Bacillus subtilis ATCC 6633 Aspergillus brasiliensis ATCC Candida albicans ATCC Environmental isolate (Staphylococcus epidermidis). Isolated from a pharmaceutical manufacturing facility in India. These organisms were selected because they represent a broad range of different microbial types and of morphological types that may occur in cleanrooms of different grades (i.e. Gram-positive coccus, Gramnegative rod, Gram-positive rod, a yeast-like fungus, and a filamentous fungus). In addition, an environmental isolate, detected from a pharmaceutical grade cleanroom was included. This organism, residential to human skin, was selected because it represented one of the most commonly recovered microorganisms from cleanrooms (with most cleanroom contamination being personnel derived) 17. The required numbers of pre-sterilised swabs were dipped into sterilised 0.9% saline tubes under a unidirectional airflow cabinet. Following this, the culture suspensions of each microorganism were prepared to provide not more than 100 CFU/0.1 ml. For the sample inoculum, a test tube containing a sterile swab stick was dipped into the 0.9% saline solution. Following this, the swab from the test tube was removed. As the swab was removed, it was squeezed against the inner walls of the test tube. This was to remove excess liquid from the swab tip. Next, 0.1 ml of not more than 100 CFU of the first microorganism was transferred onto the bud portion of the swab using a micropipette.

3 RELEASING CAPACITY OF PRE-STERILE COTTON SWABS FOR DISCHARGING SAMPLED MICROORGANISMS 123 Immediately after this, the swab stick was transferred back into the 0.9% saline solution. The procedure was repeated for each of the microorganisms. A negative control was additionally prepared during the test session. For this, a test tube containing a sterile swab stick was dipped into the 0.9% saline solution. The swab was then removed from the test tube, with swab tip squeezed against the inner walls of the test tube as it was removed. After this, 0.1 ml of a sterile 0.9% saline solution was transferred onto the bud portion of the swab using a micropipette. In doing so, the swab was treated in the same way as with the microbial, although sterile saline was used instead of a microbial. Immediately afterwards, the swab stick was transferred back into the 0.9% saline solution. Positive controls were prepared during the test session for each microorganism. Positive controls were performed by directly inoculating not more than 100 CFU of each selected organism into a tube containing 0.9% saline solution containing a swab. The samples were mixed and later membrane filtered, as per the test procedure below. Test procedure A membrane filtration unit was placed within a unidirectional airflow device. Each tube was vortex mixed for a set amount of time (initially 15 seconds, then repeated for alternative times as set out below). The vortex mixer was operating at a standard speed setting of 2500 revolutions per minute (rpm). Following vortex mixing, the swab was squeezed onto the inner walls of the test tube. After this, the swab was removed from the test tube. The contents of the test tube were then filtered through a 0.45 μm sterile filter membrane (cellulose nitrate (cellulose ester) membrane). Each filter was placed onto the surface of a pre-incubated soybean casein digest agar plate. The above exercise was performed for each test, negative control and positive control sample. The procedure was repeated by replacing the vortex period with 30 seconds, 45 second and 60 seconds. Following testing, each of the plates, including the test, negative control and positive controls, were incubated using a dual incubation regime: first at C for 3 days followed by C for 2 days (a typical order of incubation to promote the recovery of bacteria and fungi) 18. Acceptance criteria The acceptance criteria were as follows. All microbial s were to be not more than 100 CFU/0.1 ml. Although the study was an evaluation, indicative criteria were set. For this, the recovery of microorganisms in the test samples had the target of not to be less than 50% of the recovery obtained in the positive control. This was based on 50% being generally the minimum used in pharmacopoeial monographs for microbiological tests. The recovery of microorganisms obtained in the positive control should be within a factor of 2 from the calculated value of the initial concentration. No growth was to be observed on the negative control. Results The results are presented below for the different vortex mix times. For each test, all negative control results were satisfactory. All positive controls were less than 100 CFU. Table 1. Positive control recovery. Count obtained (CFU) Table 2. Test results. S. aureus % B. subtilis % P. aeruginosa % C. albicans % A. brasiliensis % Environmental isolate % Mean recovery 95%

4 124 RAVIKRISHNA SATYADA, TIM SANDLE Test set A: 15 seconds vortex Results for the samples that underwent vortex for 15 seconds are presented in Tables 1 and 2. Test set B: 30 seconds vortex Results for the samples that underwent vortex for 30 seconds are presented in Tables 3 and 4. Test set C: 45 seconds vortex Results for the samples that underwent vortex for 45 seconds are presented in Tables 5 and 6. Test set D: 60 seconds vortex Results for the samples that underwent vortex for 60 seconds are presented in Tables 7 and 8. Table 3. Positive control recovery. Count obtained (CFU) Table 4. Test results. S. aureus % B. subtilis % P. aeruginosa % C. albicans % A. brasiliensis % Environmental isolate % Mean recovery 92% Table 5. Positive control recovery. Count obtained (CFU) Table 6. Test results. S. aureus % B. subtilis % P. aeruginosa % C. albicans % A. brasiliensis % Environmental isolate % Mean recovery 91%

RELEASING CAPACITY OF PRE-STERILE COTTON SWABS FOR DISCHARGING SAMPLED MICROORGANISMS 125 Table 7. Positive control recovery. Count obtained (CFU) 33 28 28 29 31 38 Table 8. Test results. S. aureus 30 33 17 91% B.

5 RELEASING CAPACITY OF PRE-STERILE COTTON SWABS FOR DISCHARGING SAMPLED MICROORGANISMS 125 Table 7. Positive control recovery. Count obtained (CFU) Table 8. Test results. S. aureus % B. subtilis % P. aeruginosa % C. albicans % A. brasiliensis % Environmental isolate % Mean recovery 92% Summary The data indicates that satisfactory recoveries were obtained for each microorganism across all vortex times. Given that the levels obtained ranged from 81% to 111% it was decided that the data did not lend itself to further analysis. In terms of differences between the four vortex mixing times, the lowest time (15 seconds) produced the highest mean recovery. However, when dealing with small numbers, the differences across the four times are not overly significant and each passed the acceptance criteria. Based on this, and considering the lean labs philosophy, the shortest vortex time should be adopted. With the different microorganisms, each organism was adequately recovered. There were no significant variations between the organisms. Microbial recovery is shown in Figure 1. Figure 1. Microbial recovery rates over time of single-use systems and pre sterilised containers.

6 126 RAVIKRISHNA SATYADA, TIM SANDLE Discussion This study was performed to assess the releasing capacity for discharging the sampled microorganisms from presterilised cotton swabs used for the environmental sampling of surfaces. The study was performed based on the procedure outlined above. Each reference organism (0.1 ml) was spiked onto the swab bud and placed in 0.9% saline solution. The inclusion of a low-level microbial was important as the intended use of the swabs is for monitoring EU GMP Grade A/ISO 14644:2015 Class 5 environments, where any microorganisms present on surfaces would be at a very low frequency and of a very low number. The study focused on the degree of mixing required to ensure a sufficient proportion ( 50%) of microorganisms were released from the swab tip. To demonstrate this, each sample was vortexed for 15 seconds and analysed by membrane filtration technique. The procedure was repeated for each reference organism by replacing the vortexing period with 30 seconds, 45 seconds and 60 seconds. Negative and positive controls were performed and assessed against pre-set criteria. In terms of the results obtained, since the release of all the microorganisms from the cotton swabs at each vortex period are well over 50%, it is recommended that the cotton swabs, used to sample surfaces, be vortexed for not less than 15 seconds prior to testing. We also recommend that a standard vortex mix speed of 2500 rpm is used. These parameters are applicable to the type of swab tested. Other swabs may vary; hence this paper can be viewed as a case study for microbiologists to base their own swab evaluations on. Aspects not covered by the study were an evaluation of different test methods (it was taken that membrane filtering the diluent would be superior to plating the swab out onto an agar plate), and variations with the swab tip. The swab tip weave selected is becoming industry standard (that is flocked); however, there are variations to the material used to manufacture the tip. It is also possible that differences could occur with alternative diluents (or indeed with no diluent at all, although pre-wetting is generally regarded as best practice for helping to remove microorganisms that have adhered to a surface in addition to physical pressure). Possible diluents for use with swabs include Ringer s solution (an isotonic solution of sodium chloride, potassium chloride, calcium chloride and sodium bicarbonate); Dey Engley neutralising broth (which contains lecithin, Tween, sodium thiosulfate and sodium bisulfite); phosphate buffered saline; and sodium hexametaphosphate. To add to these, the storage time and temperature between sampling and plating out can also affect microbial recovery 19. These are areas for further research. Moreover, the study looked at direct s rather than the recovery of a microbial direct from a surface. With the latter, representative surfaces of pharmaceutical cleanrooms could be artificially inoculated using different environmental strains (an in vitro study) in order to review whether the type of surface has a significant effect on the recovery rates for different microorganisms (some literature, for example, suggests both surface material and surface roughness are influencing factors 20 ). While this also presents an area that is poorly researched, no evaluation of a surface environmental monitoring method can demonstrate that the method is capable of recovering all of the microorganisms attached to a surface (since the mechanisms of attachment of microorganisms to surfaces are not fully understood). Thus, we see our experimental findings as one aspect of the evaluation of swabs for the microbiological assessment of cleanroom surfaces. The additional points, together with our experimental findings, suggest that when swabs are used for environmental monitoring their suitability should be evaluated. References 1. Ljungqvist B and Reinmuller B. Some observations on environmental monitoring of cleanrooms. European Journal of Parenteral Sciences 1996;1: Sandle T. Applying spectrophotometric monitoring to risk assessments in biopharmaceutical cleanrooms. Clean Air and Containment Review 2014;Issue 20: Lewis T, Griffith C, Gallo M and Weinbern M. A modified ATP benchmark for evaluating the cleaning of some hospital environmental surfaces. Journal of Hospital Infection 2008;69: Whyte W and Eaton T. Microbiological risk assessment in pharmaceutical cleanrooms. European Journal of Parenteral and Pharmaceutical Sciences 2004;9(1): Sandle T. Environmental monitoring: a practical approach. In: Environmental Monitoring: A Comprehensive Handbook, Volume 6; Moldenhauer J, Ed. River Grove, IL, USA: PDA/DHI; 2012, pp Favero M, McDade J, Robertsen J, Hoffman R and Edwards R. Microbiological sampling of surfaces. Journal of Applied Bacteriology 1968;31: Sandle T. A study of a new type of swab for the environmental monitoring of isolators and cleanrooms (the Heipha ICR-Swab). European Journal of Parenteral and Pharmaceutical Sciences 2011;16(2): Whyte W, Carson W. and Hambraeus A. Methods for calculating the efficiency of bacterial surface sampling techniques, Journal of Hospital Infection 1989;13: Österblad M, Järvinen HJ, Lönnqvist K, Huikko S, Laippala P, Viljanto J, Arvilommi H and Huovinen P. Evaluation of a new cellulose sponge-tipped swab for microbiological sampling: a laboratory and clinical investigation. Journal of Clinical Microbiology 2003;41(5): Rose L, Jensen B, Peterson A, Banerjee SN and Arduino MJ. Swab materials and Bacillus anthracis spore recovery from nonporous surfaces. Emerging Infectious Diseases, 2004;10(6): Dalmaso G, Bini M, Paroni R and Ferrari M. Validation of the new Irradiated Nylon flocked QUANTISWAB for the quantitative recovery of micro-organisms in critical cleanroom environments. PDA Journal of Science and Technology 2008;62(3): Van Horn KG, Audette CD, Tucker KA and Sebeck D. Comparison of swab transport systems for direct release and recovery of aerobic and anaerobic bacteria. Diagnostic Microbiology and Infectious Disease 2008;62: Dalmaso G and Denoya C. Microbial control and monitoring in aseptic processing cleanrooms. Controlled Environments 1 December Available at: /01/microbial-control-and-monitoring-aseptic-processingcleanrooms (accessed 26th June 2016). 14. Sandle T. Evaluation of two different types of contact plates for microbiological environmental monitoring. European Journal of Parenteral and Pharmaceutical Sciences 2011;16(4): Scherlock O, O Connell N, Creamer E and Humphreys H. Is it really clean? 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7 RELEASING CAPACITY OF PRE-STERILE COTTON SWABS FOR DISCHARGING SAMPLED MICROORGANISMS Niskanen A and Pohja M. Comparative studies on the sampling and investigation of microbial contamination of surfaces by the contact plates and swab methods. Journal of Applied Bacteriology 1977;42: Sandle T. A review of cleanroom microflora: types, trends, and patterns, PDA Journal of Pharmaceutical Science and Technology 2011;65(4): Sandle T. Examination of the order of incubation for the recovery of bacteria and fungi from pharmaceutical cleanrooms. International Journal of Pharmaceutical Compounding 2014;18(3): Mahony JB and Chernesky MA. Effect of swab type and storage temperature on the isolation of Chlamydia trachomatis from clinical specimens. Journal of Clinical Microbiology 1985;22(5): Probst A, Facius R, Wirth R and Moissl-Eichinger C. Validation of a nylon-flocked-swab protocol for efficient recovery of bacterial spores from smooth and rough surfaces. Applied and Environmental Microbiology 2010;76(15):

á61ñ MICROBIOLOGICAL EXAMINATION OF NONSTERILE PRODUCTS: MICROBIAL ENUMERATION TESTS

USP 40 Microbiological Tests / á61ñ Microbiological Examination 1 á61ñ MICROBIOLOGICAL EXAMINATION OF NONSTERILE PRODUCTS: MICROBIAL ENUMERATION TESTS INTRODUCTION The tests described hereafter will allow