Peer Review Article | Open Access | Published 27th Sept 2020 | Submitted 28th January 2020
Study of contact plates recovery from pharmaceutical cleanroom surfaces across three-time ranges
Tim Sandle, BPL UK | EJPPS | 253, (2020) | Cite this article | Click to download pdf paper
Abstract
Viable environmental monitoring methods remain primarily culture based. One example is with the contact plate. While the method is long-established, there remain aspects that are under-researched in relation to sampling. Factors affecting surface recovery relate to microbial adhesion, the type of surface, the sampling method and the time and pressure applied. This paper examines the effect of time, when a consistent pressure is applied, in relation to microbial recovery (for the organism Staphylococcus aureus) from two surfaces common to pharmaceutical facilities: stainless-steel and vinyl. The experimental results show that surface recovery was superior for vinyl compared with stainless-steel. For both surface types, a 20 second sampling time was shown to lead to a better recovery compared to a ten second sampling time (with a 30 second sampling time not leading to a significant improvement to the microbial surface recovery).
Introduction
Environmental monitoring is always an important consideration when developing a biocontamination control strategy and it is an effective tool provided a risk-based approach has been used, covering such aspects as sample selection and for determining monitoring frequencies ¹. Through an effective environmental monitoring programme, out of limits results and variations to trend may indicate problems with cleaning and disinfection, the impact of new equipment, a drop in the standards of cleanroom operator behaviour, a problem with an air-handling system, issues with material transfer etc. The methods used for such evaluations, with the exception of emergent rapid microbiological methods, remain culture media based.
Culture-based environmental monitoring methods are long-established and carry inherent limitations in relation to their design, data-capture attributes, and culture-dependency. These methods are, conventionally, the contact plate, the active (or volumetric) air-sampler, settle plate, and swab. To these methods there are longer established alternatives, such as ATP-based swabs ² (albeit unsuitable for low-level recoveries) and newer spectrophotometric particle counters (designed to differentiate between biologic and inert airborne particles) ³. However, it remains that the culture-based methods continue to be the most widely used within pharmaceutical and healthcare facilities.
Earlier papers by this author have explored the limitation of settle plates ⁴, active air samplers ⁵ and swabs ⁶. The focus of this paper is with the contact plate, and whether method improvements – in the form of applicators designed to control time and pressure – can improve the recovery of microorganisms from surfaces or at least aid with the consistency of sampling practice. Influencing factors with surface recovery include how microorganisms adhere to cleanroom surfaces and hence the extent to which they can be recovered. This remains an under-researched area. With the study, an applicator that controls both time and pressure was used. While the pressure was fixed, the factor of time was varied.
The research question examined was – does the time that a contact plate remains in contact with a surface affect microbial recovery? This was assessed using two different types of surfaces – stainless steel and vinyl. The results showed that better recovery was obtained from vinyl compared with stainless steel and that higher recoveries are achieved with a 20 second contact time compared with a ten second contact time. The data also informs as to the typical proportion of microbial numbers that a contact plate can consistently recover from a surface. This proportion was found to be relatively low, at no more than 25% (with this level of recovery compared against multiple plates used on the same area of inoculated surface).
Limitations of culture-based monitoring methods
With any culture based environmental monitoring method there will be limitations based on the type and formulation of the culture medium selected. There is no ‘universal culture medium’ and there is no set of incubation conditions that can recover all cleanroom microbial isolates, with respect to temperature and time ⁷. This is both a product of the regime selected against a given population of organisms and due to the nature of non-culturability, which is both medium dependent and a factor of many cleanroom isolates being sub lethally damaged or under conditions of stress which impart a fragility that makes recovery on standard media very difficult (the oft termed ‘viable but non-culturable’ organisms).
In the context of these limitations, microbiologists should set out to ensure that:
The medium and incubation conditions selected have a reasonable chance of detecting the main organisms associated with the cleanroom environment (these will be those organisms associated with people, and the Human Microbiome Project has provided valuable supporting data for this) ⁸.
Extraneous factors that could affect microbial recovery are addressed (such as the addition of a suitable neutralizer to the culture medium for the monitoring of surfaces or the gloved hands of operators) ⁹,¹⁰.
Developing methods and procedures that ensure consistency of practice.
With the last issue, this is one reason why trend analysis is important: for any given monitoring session the optimal recovery may not always be obtained, but by operating the best methods possible and consistently, the microbiologist has a better chance of examining one set of results against another for understanding the formation of an upward trend, and with then taking action at the most appropriate time.
Contact plates
Contact plates are the primary method of choice for quantitatively sampling flat, impervious surfaces and floors for the presence of microorganisms, generally showing superiority over swabbing ¹¹,¹²,¹³. Swabs are best reserved for narrow or non-flat surfaces. However, as with any of the ‘classic’ environmental monitoring methods there are concerns with recovery (in this case, overcoming surface attachment and loss of viability) and readability, such as when clumping or merging of colonies occurs ¹⁴, which can affect the reliability of plate counting ¹⁵. In addition to contact plates and swabs, there are alternatives for surface monitoring, such as agar slides ¹⁶, and with nitrocellulose membranes or chemo luminescence technologies, but these have not been widely adopted as part of environmental monitoring programmes ¹⁷.
The contact plate is filled so that the media forms a dome, and the plate diameter has been generally standardized at 55 millimetres to ensure that a 25cm2 area of the surface to be sampled. The current contact plate is based on the original (and trademarked) RODAC plate, an acronym for ‘replicate organism detection and counting’ (first proposed by Hall and Hartnett ¹⁸), which was developed during the 1960s ¹⁹. Contact plates are produced by different culture media manufacturers to approximate the original RODAC design; however, there will be differences in the preparation of the agar, especially with the way a plate is filled and consequently the degree to which the dome forms of the plate. There will also be variations with the formulation of the culture media ²⁰.
To make the act of sampling a surface using a contact plate more consistent, several companies manufacture contact plate holding devices (or applicators). A contact plate applicator ensures that a controlled pressure is applied. This means that each operator undertaking sampling applies the agar dome to a surface at the same pressure, which is a factor of a weight ensuring that a consistent force is applied, as per:
P = F/A
Where:
P is the pressure
F is the applied force
A is the surface area where the force is applied
Such devices also control the length of time, ensuring that sampling time is consistent for each sample taken. While applicator devices can create more consistent conditions, this does not mean that microorganisms are necessarily recovered from a given surface.
Factors affecting surface recovery
The majority of microorganisms in the biosphere are attached to surfaces and microorganisms prefer this sessile condition to the free-floating or planktonic state ²¹. When bacteria attach to surfaces, they undergo metabolic changes with such changes becoming more complex if a biofilm forms. These changes include swarming and surface sensing is a precursor to this adaptive behaviour, whereby contact between cells and surfaces programmes morphological changes, such as cooperative behaviour, rapid community growth, and migration of communities. Microbial attachment to horizontal surfaces stimulates bacterial growth (particularly in nutrient-poor environments) as organic material suspended in liquid settles, is deposited on surfaces, and this increases the local concentration of nutrients ²². In addition, some bacteria obtain necessary metabolites and co-factors directly from the surfaces to which they adhere.
With surface attachment, initial attachment is reversible and occurs rapidly (in less than one minute). This process involves hydrodynamic and electrostatic interactions, whereby the adhesive force between bacteria and surfaces increases rapidly. This is largely based on physicochemical effects (such as the loss of interfacial water and structural changes in surface molecules), rather than biological effects. This phase is followed by attachment that is irreversible, and this occurs on a time scale of several hours ²³. By irreversible this means a state when microbes can no longer move perpendicularly away from the surface. This process involves van der Waals interactions between the hydrophobic region of the outer cell wall and the surface, and there are biological effects which will continue, such as changes to proteins which aid the transition from reversible to irreversible cell attachment. The process has been described in greater detail by Sandle ²⁴.
Evidence suggests that the ability to colonize a surface helps to promote bacterial survival within a cleanroom ²⁵. Increased knowledge of this process is leading different strands of research to either add additives to materials that can exert antimicrobial effects (such as silver) or to alter surfaces to repel bacteria, such as the use of anodization to create nanoscale pores that change the electrical charge and surface energy of a metal surface, which in turn exerts a repulsive force on bacterial cells and prevents attachment (and also biofilm formation) ²⁶. Alternatively, adding a titanium coating to stainless steel discourages the retention of many bacteria.
In terms of removing microorganisms from surfaces, for the purposes of environmental monitoring, there are different factors which influence surface recovery. One factor is the types of surface (variation of attachment substrata), for different surfaces lead to different recoveries, as assessed in different studies over multiple impressions ²⁷. Bacteria are able to attach to a wide variety of different materials, including glass, aluminium, stainless steel, various organic polymers, and fluorinated materials ²⁸. A related factor is the age of the surface and to the extent that the surface is worn, pitted or damaged. Another factor is the way that microorganisms are attached to the surface, which is itself varied according to soiling (organic or inorganic matter), organism type and physicochemical interactions. Bacterial adhesion to different types of surfaces will also vary. This is affected by surface charge, wettability, topography (such as the ‘roughness’ of the surface) and hydrophobicity ²⁹. With surface roughness, the smoother the surface the less likely some types of microbial cells are to stick. There are three categories of surface roughness, termed as macro (Ra ~ 10 μm), micro (Ra ~ 1 μm) and nano roughness (Ra ~ 0,2 μm) ³⁰. A rougher surface has more nooks and crannies where microbes can get lodged, so lower Ra values are considered more hygienic. With wettability, surfaces with moderate wettability are more able to bind bacteria or cells, as compared with extremely hydrophobic or hydrophilic surfaces ³¹.
A further factor relates to the types of microorganisms. Bacillus spores, for example, adhere as monolayers on many kinds of surfaces, hydrophobic spores of B. cereus being the most adhesive. A related variable will be whether organisms are present as one species or as a community of different species (the latter becoming more likely in lower grade cleanrooms).
In relation to the complexities of adhesion, removing microorganisms from a surface will be affected by the method deployed. Where contact plates are used, as with this study, factors affecting recovery will include variations with manufacturers of ready-to-use culture media items, the surface composition of the agar (notably the physical dimensions of the dome structure) ³², pressure (which is proportionate to the weight applied), and the time of contact, plus variances with incubation parameters as previously discussed. Other factors that are difficult to control include the distribution of organisms, which can lead to colonies merging if they are in close proximity to each other ³³. In using an applicator with a standardized weight, this study does not look at pressure but at time.
Although there are many factors affecting recovery, and each is of importance, the microbiologist still needs to undertake surface sampling, both to assess the potential risk of product contamination and to conform with regulatory expectations. Hence, the factors cannot be used as mitigations for inaction, but rather they inform the microbiologist as to the limitations of the method and provide some guidance as to how recovery might be improved. The purpose of the study described in this paper is to measure contact plate sampling limitations and to explore the extent that contact time is a factor in microbial recovery.
Method
The method evaluated was one of the two standard methods used for recovering microorganisms from cleanroom surfaces – the contact plate, containing tryptone soya agar. Tryptone Soya Agar (TSA) is a non-selective agar used for the isolation and enumeration of a range or organisms, aerobic and anaerobic bacteria, yeast and fungi. It is presented as a contact plate with neutraliser N◦4 plates. The formulation of the medium was:
Tryptone soya agar (Oxoid – CM1118)
Agar No3 (Oxoid LP0013)
Neutraliser No. 4:
Lecithin (Sigma P5638)
Tween 80 (Sigma P1754)
Sodium thiosulphate (Sigma S8503)
L-histidine (Sigma H8000)
Hydrochloric acid (37%)
1:9 dilution (Prolabo, 30024.290) and demineralized water
Two surfaces were examined for the study: Stainless steel Grade 316 and polyvinyl chloride (2mm thick sheet vinyl cladding). For the sampling, a contact plate applicator was used. This device holds the plate securely and enables a controlled weight to be applied (500 g, with the applicator used). The time can be set by the user. For this study three time points were selected: 10, 20 and 30 seconds. It was reasoned that ten seconds would be the minimum to ensure recovery of a bacterium from a surface and that a maximum of 30 seconds was in keeping with the work of a microbiologist who may need to monitor several samples across multiple cleanrooms (in that the task could not be unnecessarily long, otherwise this would begin to impact upon operational efficiencies).
The surfaces were challenged with a selected microorganism, and run in triplicate, on the two different surface types. With each of the triplicate tests, each inoculated area was sampled with 10 contact plates repeatedly for one of the specified contact times. Although for environmental monitoring in practice only one sample would be taken, the use of ten replicates was intended show how far away from maximum potential recovery one sample would be, drawing on a method proposed by Tidswell and colleagues to control time, albeit under circumstances where pressure was not controlled ³⁴.
The microorganism selected for the study was Staphylococcus aureus, which was obtained from a recognizable culture collection (ATCC 6538). Only one microorganism was selected for testing due to the nature of the study which was to evaluate the contact plates themselves and not their ability to sample multiple microorganism types, although this is recognised as a limitation that requires further work, as noted in the discussion section. The microorganism selected was from a typical genus found in cleanrooms, and of a species which can be recovered depending upon the extent that operators are host to this particular pathogen. The organism itself was also one of those recommended in the European Pharmacopeia for the evaluation of samples against the Microbial Limits Test.
Testing was conducted inside a unidirectional airflow cabinet and used triplicate stainless steel and vinyl test surfaces. Each surface was inoculated with a ≤ 100 CFU microbial suspension and spread across demarcated areas of a surface area of 25cm2. The suspension was then allowed to dry for a period of 40 minutes (a previously established drying time for this microorganism on surfaces). Following the 40-minute drying period, 10 replicate surface samples were taken of each inoculated 25cm2 marked area using TSA contact plates with a neutralizing agent. Separate TSA plates were inoculated, in duplicate. with a ≤ 100 CFU microbial suspension (inoculating 10μl of a diluted bacterial suspension), using the spread plate technique, to serve as positive controls.
The plates were incubated at 30-35◦C for 3 days. Following incubation, the plates were counted using both an angle poise lamp (white light source) and a colony counter equipped with a magnifying glass.
Results
The data for each replicate was averaged (by taking the mean) and presented for each time point. The positive control plates recorded a mean CFU of 64. The positive controls established the theoretical level of microorganism on the surface.
Ten second sampling time
Table 1 shows that the recoveries from vinyl were greater compared with stainless steel. After ten replicate plates, a 63% recovery was obtained for vinyl surfaces compared with 39% from a stainless-steel surface. From the first contact, the recoveries were 13% from stainless steel and 28% from vinyl.
The cumulative percentage recovery, for both surface types is displayed in Figure 1:
Ten second sampling time
Table 1 shows that the recoveries from vinyl were greater compared with stainless steel. After ten replicate plates, a 63% recovery was obtained for vinyl surfaces compared with 39% from a stainless-steel surface. From the first contact, the recoveries were 13% from stainless steel and 28% from vinyl.
The cumulative percentage recovery, for both surface types is displayed in Figure 1:
Thirty second sampling time
Table 3 shows that the recoveries from vinyl were and stainless steel were again relatively close, and again this contrasted with the ten second sampling time. However, the variations between the 20 and 30 second contact times were not great. After ten replicate plates, a 59% recovery was obtained for vinyl surfaces compared with 52% from a stainless-steel surface. From the first contact, the recoveries were 17% from stainless steel and 30% from vinyl.
Discussion
With single contact plates the recovery of viable microorganism was poor, and there were differences with the two surfaces examined. Notably, recoveries were higher for vinyl compared with stainless steel, this was wider at the ten second sampling time although this narrowed as the sampling time was extended.
In terms of the limitation of the method, the percentage of recovery from the contact plates was relatively low based on the first plate applied to a surface, which matches the way a sample would be taken within the cleanroom environment. For a ten second sampling time the recovery was 13% (stainless steel) and 28% vinyl (mean 20%); for a 20 second sampling time the recoveries were 19% (stainless steel) and 31% vinyl (mean 25%). With the 30 second contact time, recoveries were 17% (stainless steel) and 30% vinyl (mean 24%).
The data shows recoveries improved between 10 and 20 seconds, albeit by a small amount from one plate, and to a greater extent when replicate counting was taken into consideration (refer to figures 4 and 5).
Extending the sample time to 30 seconds did not make a large difference and it can be argued that extending the time from 20 to 30 seconds does not add value, however, increasing the contact time from 10 to 20 seconds appears to add some value in terms of improving the surface count estimate. This is shown by comparing figure 5 to figure 6.
With the test control, the results showed that the microbial population on the test surface is viable after a drying period of 42 minutes in a unidirectional flow safety cabinet, and that desiccation has no effect on micro-organisms during this test procedure.
Based on these results, improvements can be made with sampling time, although microbiologists will still need to be aware that recoveries remain underestimations even with those organisms that can theoretically be recovered using the culture medium, time and temperature combinations used in this study. The findings in this study match some of the concerns about method limitations from literature. A study by Pinto and colleagues, which compared three different brands of contact plates using a similar applicator to the one used in this study (although only for ten seconds and against a stainless-steel surface), showed recoveries for two species of Staphylococcus ranging from 23 to 56% (mean – 38%). There were variations with the recovery of the two species of Staphylococci (S. aureus gave a better recovery than S. epidermidis); this was consistent with the findings of other surface recovery research conducted by Scott and associates ³⁵, and it is of interest given that both species of Staphylococci will be found in cleanrooms (albeit with S. epidermidis more likely to be recovered in higher numbers given this organism’s greater ubiquity on the outer layers of the skin and only a minority of the population being carriers of S. aureus) ³⁶.
In assessing the data, the low recovery may be due to the inability of the dried microorganism to transfer to the plate, rather than the inability of the plate to grow the microorganism. While this is interesting, organisms in a drier state will be more representative of what is likely to be found within a cleanroom. Wetness might help in improving recovery. Furthermore, although a common cleanroom bacterium was used this organism may or may not be representative of other typical cleanroom-associated organisms. A more extensive assessment involving other organisms would be required to draw a wider inference. The author has undertaken an assessment of organism recovery from the two surfaces, but not as yet as a time-based study ³⁷.
With the differences between surface types, the low total percentage of the microbial population recovered by the stainless-steel samples tested with a 10 second contact time may relate to the rougher surface of the stainless steel when compared with the vinyl surface, which causes difficulty in sampling microorganisms from the surface due to adhesion to microscopic pits in the steel. One observation made during the test procedure was the increase in moisture of the sampling surface when a 20 second contact time was used, which was assessed by visual examination. The level of moisture did not vary between the 20 second and 30 second sampling times. The poor recovery on the previous 10 second recovery test recovery was could be due to the inability of the dried microorganism to transfer to the plate, the moisture on the stainless-steel surface during the 20 second recovery test may have improved recovery
Underestimating the microbial contamination of surfaces might have serious consequences on the quality assurance of aseptically prepared pharmaceutical products. Another part of the study looked at recoveries when consecutive samples were taken. Based on the increase to the microbial count as more samples were taken it might be that under specific circumstances consecutive replicate contact plate sampling may represent a more appropriate means of evaluating surface-borne bioburden, perhaps under conditions where high bioburden is likely to be present. However, once the method has been standardized in terms of controlling weight and time has been optimized (balancing what is practical against what provides a greater recovery), microbiologists must remain cognizant that the levels of organisms recovered from a cleanroom surface are likely to be an underestimation of the organisms attached to the surface.
References
01. Sandle, T. (2012). Environmental Monitoring: a practical approach. In Moldenhauer, J. Environmental Monitoring: a comprehensive handbook, Volume 6, PDA/DHI: River Grove, USA, pp29-54
02. Amodio, E. and Dino, C. (2014) Use of ATP bioluminescence for assessing the cleanliness of hospital surfaces: a review of the published literature (1990-2012), J Infect Public Health. 7(2):92-8
03. Sandle, T. (2014) Applying spectrophotometric monitoring to risk assessments in biopharmaceutical cleanrooms, Clean Air and Containment Review, Issue 20, pp22-25
04. Sandle, T. (2015) Settle plate exposure under unidirectional airflow and the effect of weight loss upon microbial growth, European Journal of Parenteral & Pharmaceutical Sciences; 20(2): 45-50
05. Sandle, T. (2010) Selection of active air samplers, European Journal of Parenteral and Pharmaceutical Sciences, 15 (4): 119-124
06. Sandle, T. (2011). 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, 16 (2): 42-48
07. Sandle, T. (2014) Examination of the Order of Incubation for the Recovery of Bacteria and Fungi from Pharmaceutical Cleanrooms, International Journal of Pharmaceutical Compounding, 18 (3): 242 – 247
08. Proctor, L. M. (2016) The National Institutes of Health Human Microbiome Project, Semin Fetal Neonatal Med.;21(6):368-372
09. Schiemann, D.A and Chatfield, D.E. (1976) Evaluation of neutralizers in Rodac media for microbial recovery from disinfected floors, J Environ Health. 38(6):401-4
10. Sutton, S.V. W., Proud, D. W., Rachui, S. and Brannan, D. K. (2002) Validation of Microbial Recovery From Disinfectants, PDA Journal of Pharmaceutical Science and Technology, 56 (5): 255-266
11. Lemmen, S.W., Häfner, H., Zolldann, D., Amedick, G. and Lütticken, R. (2001) Comparison of two sampling methods for the detection of gram-positive and gram-negative bacteria in the environment: moistened swabs versus Rodac plates, Int J Hyg Environ Health. 203(3):245-8
12. Niskanen, A. and Pohja, M.S. (1977) Comparative studies on the sampling and investigation of microbial contamination of surfaces by the contact plate and swab methods. J Appl Bacteriol 42, 53–63
13. Lemmen, S.W., Häfner, H., Zolldann, D., Amedick, G. and Lütticken, R. (2001) Comparison of two sampling methods for the detection of Gram‐positive and Gram‐negative bacteria in the environment: moistened swabs versus Rodac plates. Int J Hyg Environ Health 203: 245–248
14. Favero, M.S., McDade, J.J., Robertsen, J.A., Hoffman, R.K. and Edwards, R.W. (1968) Microbiological sampling of surfaces. J Appl Bacteriol 31, 336–343
15. Tidswell, E. C. and Sandle, T. (2017) Microbiological Test Data - Assuring Data Integrity, PDA J Pharm Sci Technol; 72 (1): 2-14 doi:10.5731/pdajpst.2017.008151
16. Griffiths, W. E., Contact slides for use in environmental hygiene studies. Environ. Health. 87:36-37, 1978
17. Poletti, L., Pasquarella, C., Pitzurra, M. and Savino, A. (1999) Comparative efficiency of nitrocellulose membranes versus RODAC plates in microbial sampling on surfaces, J Hosp Infect. 41(3):195-201
18. Hall L.B and and M.J. Hartnett (1964) Measurements of Bacteria contamination on surfaces in Hospitals, Public Health Rep 79 (11):1021-1024
19. Brewer, J. A. and Turner, A. G. (1973) Replicating RODAC plates for identifying and enumerating bacterial contamination, Health Lab. Sci. 10: 195–202.
20. Sandle, T. (2011): Evaluation of two different types of contact plates for microbiological environmental monitoring, European Journal of Parenteral and Pharmaceutical Sciences, 16 (4): 116-120
21. Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, Lappin-Scott HM. (1995) Microbial biofilms Annu. Rev. Microbiol. 1995;49:711–745
22. ZoBell CE. (1943) The Effect of Solid Surfaces upon Bacterial Activity; J Bacteriol, 46:39–56
23. Renner LD, Weibel DB. (2011) Physicochemical regulation of biofilm formation, MRS Bull. 36:347–355
24. Sandle, T. (2013). Bacterial Adhesion: an Introduction, Journal of Validation Technology, 19 (2): 1-10 - on-line: http://www.ivtnetwork.com/article/bacterial-adhesion-introduction
25. La Duc, M.T., Dekas, A., Osman, S., Moissl, C., Newcombe, D. & Venkateswaran, K. (2007a). Isolation and characterization of bacteria capable of tolerating the extreme conditions of clean room environments. Appl. Environ. Microbiol. 73, 2600–2611
26. Guoping Feng, Yifan Cheng, Shu-Yi Wang, Lillian C. Hsu, Yazmin Feliz, Diana A. Borca-Tasciuc, Randy W. Worobo & Carmen I. Moraru (2014) Alumina surfaces with nanoscale topography reduce attachment and biofilm formation by Escherichia coli and Listeria spp., Biofouling, 30:10, 1253-1268
27. Werner, H-P., Swinke, U. and Werner, G. (1977) Development of a new test method for surface disinfection procedures. III. The impression method: influence of the test surface material and the types of microorganisms on the recovery rate, Zentralbl Bakteriol Orig B. 165(1):20-42
28. Banerjee I, Pangule RC, Kane RS. (2011) Antifouling coatings: recent developments in the design of surfaces that prevent fouling by proteins, bacteria, and marine organisms, Adv. Mater.;23:690–718
29. Flint, S.H., Brooksa, J.D. and Bremer, P.J. (2000) Properties of the stainless steel substrate, influencing the adhesion of thermo‐resistant streptococci. J Food Eng 43, 235–242
30. M. G. Donoso, A. Méndez-Vilas, J. M. Bruque, and M. L. González-Martin (2007) “On the relationship between common amplitude surface roughness parameters and surface area: Implications for the study of cell-material interactions,” International Biodeterioration & Biodegradation, 59 (3): 245–251
31. X. Zhang, L. Wang and E. Levänen, (2013) Superhydrophobic Surfaces for the Reduction of Bacterial Adhesion, RSC Adv., 3: 12003–12020
32. Bruch. M.K. and Smith, F.W. (1968) Improved method for pouring Rodac plates, Appl Microbiol. 16(9):1427-8
33. Vesley, D., Keenan, K. M., and Halbert, M. M. (1966) Effect of Time and Temperature in Assessing Microbial Contamination on Flat Surfaces, Appl Microbiol. 14(2): 203–205
34. Tidswell, E.C., Bellinger, M., McCullough, D. and Alexander, A. (2006) Consecutive replicate contact plate sampling assists investigative characterisation of surface-borne bioburden, European Journal of Parenteral Science, 10 (4): 93-96
35. Scott, E., Bloomfield, S.F. and Barlow, C.G. (1984) A comparison of contact plate and calcium alginate swab techniques for quantitative assessment of bacteriological contamination of environmental surfaces. J Appl Bacteriol 56, 317–320
36. Sullivan, S. B., Kamath, S., McConville, T. H., Gray, B. T., Franklin D. Lowy, F. D., Gordon, P. G., and Uhlemann, A-C. (2016) Staphylococcus epidermidis Protection Against Staphylococcus aureus Colonization in People Living With Human Immunodeficiency Virus in an Inner-City Outpatient Population: A Cross-Sectional Study, Open Forum Infect Dis. 3(4): ofw234
37. Sandle, T. (2019) Assessment of the recovery of different bacteria from two cleanroom surface materials, Chimica oggi (Chemistry Today), 37(5): 31-33
Author Information
Corresponding Author: Tim Sandle
Bio Products Laboratory
England
Email: timsandle@btinternet.com
Comentarios