Surface-attached cells, biofilms and biocide 
susceptibility: Implications for hospital 
cleaning and disinfection
Authors: J.A. Otter, K. Vickery, J.T. Walker, E. 
deLancey Pulcini, P. Stoodley, S.D. Goldenberg, J.A.G. 
Salkeld, J. Chewins, S. Yezli, & J.D. Edgeworth
NOTICE: this is the author’s version of a work that was accepted for publication in Journal of 
Hospital Infection. Changes resulting from the publishing process, such as peer review, editing, 
corrections, structural formatting, and other quality control mechanisms may not be reflected in 
this document. Changes may have been made to this work since it was submitted for 
publication. A definitive version was subsequently published in Journal of Hospital Infection, 
VOL# 89, ISSUE# 1, (January 2015) DOI# 10.1016/j.jhin.2014.09.008
Otter JA, Vickery K, Walker JT, de Lancey Pulcini E, Stoodley P, Goldenberg SD, Salkeld JA, 
Chewins J, Yezli S, Edgeworth JD, "Surface-attached cells, biofilms and biocide susceptibility: 
Implications for hospital cleaning and disinfection," Journal of Hospital Infection. January 2015 
89(1):16–27.
Made available through Montana State University’s ScholarWorks 
scholarworks.montana.edu 
Surface-attached cells, biofilms and biocide 
susceptibility: implications for hospital 
cl
ery 
.G.
Diagn
t, Lond
edicin
on Do
Monta
on and
ibolog
reduce microbial attachment and improve microbial detachment, and methods to augment the 
activity of biocides against surface-attached microbes such as bacteriophages and antimicrobial 
peptides. Future strategies to address environmental contamination on hospital surfaces should 
consider the presence of microbes attached to surfaces, including biofilms.Microbes tend to attach to available surfaces and readily form biofilms, which is prob-lematic in 
healthcare settings. Biofilms are traditionally associated with wet or damp surfaces such as 
indwelling medical devices and tubing on medical equipment. However, microbes can survive for 
extended periods in a desiccated state on dry hospital surfaces, and biofilms have recently been 
discovered on dry hospital surfaces. Microbes attached to surfaces and in biofilms are less 
susceptible to biocides, antibiotics and physical stress. Thus, surface attachment and/or biofilm 
formation may explain how vegetative bacteria can survive on surfaces for weeks to months (or 
more), interfere with attempts to recover microbes through environmental sampling, and provide a 
mixed bacterial population for the horizontal transfer of resistance genes. The capacity of existing 
detergent formula-tions and disinfectants to disrupt biofilms may have an important and previously 
unrec-ognized role in determining their effectiveness in the field, which should be reflected in 
testing standards. There is a need for further research to elucidate the nature and physiology of 
microbes on dry hospital surfaces, specifically the prevalence and compo-sition of biofilms. This 
will inform new approaches to hospital cleaning and disinfection, including novel surfaces that 
SummaryJ.A. Otter a, b, *, K. Vick
S.D. Goldenberg a, J.A
aCentre for Clinical Infection and
St. Thomas’ NHS Foundation Trus
bBioquell (UK) Ltd., Andover, UK
cAustralian School of Advanced M
d Public Health England MSD, Port
eCenter for Biofilm Engineering,
fDepartments of Microbial Infecti
gNational Centre for Advanced Treaning and disinfection
c, J.T. Walker d, E. deLancey Pulcini e, P. Stoodley f, g,
 Salkeld b, J. Chewins b, S. Yezli b, J.D. Edgeworth a
ostics Research, Department of Infectious Diseases, King’s College London and Guy’s and
on, UK
e, Macquarie University, Nth Ryde, Australia
wn, Salisbury, UK
na State University, Bozeman, MT, USA
Immunity and Orthopedics, Ohio State University, Columbus, OH, USA
y, University of Southampton, Southampton, UK
Introduction
Microbes tend to attach to available surfaces and form
biofilms readily.1e3 Biofilms are problematic in healthcare
settings, where they are thought to be involved in 65% of
nosocomial infections, and are usually reported in relation to
indwelling medical devices and prostheses, water lines and
tubing on endoscopes, and on wounds.1e3 In these settings,
biofilm persistence can be prolonged, periodically ‘sloughing
off’ and releasing planktonic bacteria that may act as a source
of infection. Biofilms are a common problem on liquidehard
surface interfaces, and in areas of a hospital that are usually
wet or damp, such as taps and sink drains.4 The recent prob-
lems caused by Pseudomonas aeruginosa in water supplied to
intensive care units, which resulted in changes to UK national
guidance, illustrates this problem.4
A biofilm is a community of micro-organisms attached to a
substrate producing extracellular polymeric substances (EPS)
and exhibiting an altered phenotype compared with corre-
sponding planktonic cells, especially regarding growth, gene
transcription, protein production and intercellular inter-
action.1e3,5,6 Biofilms comprising various micro-organisms,
including bacteria, viruses, fungi and other micro-organisms,
can form on almost any biological or inanimate surface, and
have been identified in various industrial and clinical set-
1,7
survival is not surprising for the metabolically inert bacterial
endospores, survival of some vegetative bacteria that is
measured in years rather than days challenges our under-
standing of bacterial physiology.10,14 The structural and phys-
iological state of microbes dried on to hospital surfaces has not
been studied in detail, but it seems likely that bacteria attach
to surfaces to some degree, and may form biofilms. Indeed, a
recent study from Australia by Vickery et al.15 ‘destructively
sampled’ (i.e. cut the materials out of the hospital environ-
ment and undertook laboratory analysis) several hospital sur-
faces after cleaning and bleach disinfection. Scanning electron
microscopy was used to examine the surfaces for biofilms,
which were identified on five of six surfaces. Furthermore,
viable meticillin-resistant Staphylococcus aureus (MRSA) was
identified in the biofilm on three of the surfaces.
This article will review in-vitro studies that explore the
structure, physiology and biocide susceptibility of microbes
dried on to hard surfaces in the context of surface attachment
and biofilm establishment, and discuss the potential implica-
tions for hospital cleaning and disinfection.16
Search strategy
Pubmed was searched with no date restrictions using the
search terms ‘biofilm and biocide’, ‘biofilm and reduced sus-
ic c
vel
tura
d b
r miillustrates EPS. The biofilm development and maturation process is a complex step-wise process, simplified here as a single step. Whilst
the reduced biocide susceptibility associated with surface attachment and biofilms will be determined by a number of factors, not leasttings. Not all microbes attached to surfaces meet the defi-
nition of a biofilm, and the transition from a planktonic culture
through surface attachment to an established biofilm is likely
to be a continuum rather than a stepwise process (Figure 1).1e3
Microbes including bacterial spores, vegetative bacteria,
fungi and viruses can also survive on dry surfaces for extended
periods.8e10 Contaminated environmental surfaces are an
increasingly recognized reservoir in the transmission of certain
healthcare-associated pathogens.11e13 Whilst this extended
Plankton
Surface 
attachment
Biofilm de
and ma
Up to 10x less 
susceptible
Figure 1. Schematic of surface attachment, biofilm formation an
faces, development and maturation of biofilms, and implications fothe biocide, microbe and testing conditions, bacteria in mature bio
surfaces, often by several orders of magnitude.18e20ceptibility’, ‘biofilm and [MRSA, VRE, C. difficile, Acineto-
bacter, E. coli, Pseudomonas]’ and ‘susceptibility planktonic
biofilm biocide’ (see Figure 1). The reference lists of articles
identified via the Pubmed searches were hand searched to
identify other relevant literature.
Resistance and reduced susceptibility
Biofilms constitute a protected mode of growth, allowing
bacteria to survive in hostile environments and conferring
ells
Mature 
biofilm
opment 
tion
Up to 1000x less 
susceptible
iocide susceptibility. This illustrates bacterial attachment to sur-
crobial susceptibility. The grey shading around the mature biofilms
2,3
films are consistently less susceptible than biofilms attached to
reduced susceptibility to dehydration, phagocytosis, metal
toxicity, acid exposure, antibiotics and biocides.1,7,17 Microbes
attached to surfaces that have not formed an established
biofilm appear to represent an intermediate step, with reduced
susceptibility to biocides compared with planktonic cells, but
increased susceptibility relative to biofilms (Figure 1).18e20
Mechanisms of reduced susceptibility
Causes of reduced susceptibility to antimicrobial agents in
biofilms are multi-factorial, including reduced penetration
(particularly due to changes in cell density and the production
of EPS), slow growth (and subsequent reduced metabolism of
antimicrobial agents), modulation of the stress response and
other metabolic processes, and changes in quorum
sensing.5,21e23 It seems likely that these mechanisms also
explain reduced biocide susceptibility in surface-attached cells
that have not yet formed biofilms.
Biocide susceptibility
Many studies have evaluated the impact of established
biofilms on biocide susceptibility, and fewer studies have
evaluated the susceptibility of surface-attached cells that have
not yet formed established biofilms (Figure 1). Table I sum-
marizes studies that have investigated organisms and biocides
relevant to disinfection in healthcare settings that include data
comparing susceptibility in planktonic culture with surface-
attached cells and/or biofilms. Studies have evaluated a
range of organisms (both alone and in combination), various
suspending media, and several methods of attaching cells to
surfaces and producing biofilms on different substrates; all of
these factors are likely to influence biocide susceptibility. One
important factor is the maturation of the microbes tested,
which ranges from cells attached to surfaces for hours to
samples extracted from continuously fed biofilm reactors that
are weeks old.24,25 Furthermore, few studies have controlled
for cell density in attached cells or biofilms compared with
planktonic culture.26 Thus, although several studies have sug-
gested that cell density alone does not explain the reduced
susceptibility of biofilms to biocides, it is difficult to be certain
of the impact of the biofilm phenotype independent of cell
density in many studies.20,27e29 A number of different ap-
proaches have been taken to quantify growth, including both
direct microbial culture and indirect measures, such as live/
dead viability assays.20,30e32 Finally, different approaches to
compare susceptibility in planktonic culture and biofilms have
included measuring the amount of biocide required to inhibit
growth [minimum inhibitory concentration (MIC)] or kill cells
[minimum bactericidal concentration (MBC)],7,18,20,33 or
measuring the survival time at a given concentration of
biocide;20,32,34,35 this makes comparison of studies difficult.
Notwithstanding difficulties in comparing studies, the phase
of the surface-attached cells influences biocide susceptibility.
In general, bacteria in planktonic culture are more susceptible
than attached cells, which are, in turn, more susceptible than
established biofilms (Figure 1).18e20 Meanwhile, detached
biofilm cells revert to the susceptible phenotypic state.19,36,37
Similarly, growth phase affects biocide susceptibility of
planktonic culture.38e40 Reduced susceptibility in surface-
attached cells ranges widely from two-fold to >1000-
fold.26,41 For example, clinical isolates of MRSA and
P. aeruginosa were grown as biofilms on discs of common ma-
terials in the hospital environment, and treated with threecommonly used hospital biocides: benzalkonium chloride (1%
w/v), chlorhexidine gluconate (4% w/v) and triclosan (1% w/
v).7 The MBCs of all biocides for planktonic cultures of both
organisms were considerably less than the concentrations
recommended for use by the manufacturer. However, when
isolates were grown as biofilms, the biocides were ineffective
at killing bacteria at the concentrations recommended for use.
The MBCs of all three biocides were found to be 10e1000-fold
higher than the same isolates grown in planktonic culture for
MRSA and P. aeruginosa. Following biocide treatment, up to
11% of cells in MRSA biofilms survived, and up to 80% of cells in
P. aeruginosa biofilms survived. Another study evaluated the
susceptibility of four Candida spp. and two Escherichia coli
strains to sodium hypochlorite, ethanol, hydrogen peroxide and
iodine.20 Strains were tested in planktonic culture, as attached
cells and as biofilms in microtitre plates. Whilst susceptibility
varied by organism and biocide, biofilms were less susceptible
than attached cells, which were less susceptible than plank-
tonic cells. MICs for biofilms were up to >10-fold higher for 5-
min and 24-h exposures compared with planktonic cells. These
studies suggest that although biocides may be effective against
planktonic populations of bacteria, some biocides currently
used in hospitals may be ineffective against nosocomial path-
ogens when attached to surfaces or in biofilms, and thus fail to
control this reservoir for hospital-acquired infection.7
However, whilst surface-attached microbes and biofilms are
generally less susceptible to biocides than bacteria in plank-
tonic culture, the degree of reduced susceptibility is not al-
ways this stark. For example, a study reported no difference
between planktonic culture and biofilms of Klebsiella pneu-
moniae exposed to sodium hypochlorite and monochlor-
amine.42 Other studies have not identified reduced
susceptibility for all biocides or organisms tested.20,24,43,44 It is
difficult to determine the relative importance of organism,
biocide and testing conditions in these studies that found little
or no reduced biocide susceptibility associated with biofilms.
The composition of the biofilm also influences susceptibility.
For example, high-nutrient, high-density biofilms are less sus-
ceptible to biocides than low-nutrient, low-density bio-
films.18,31,45 This seems particularly important in the context of
biofilms thatmaybepresent onhospital surfaces,which are likely
to be low-nutrient, low-density biofilms in most cases. However,
gross contamination with body fluids could provide an environ-
ment in which high-nutrient, high-density biofilms could form on
hospital surfaces. Indeed, three-quarters of thebiofilms reported
byVickeryetal.had very thick EPSdespite having a lowdensity of
microbes inmost cases, perhaps in response todesiccation.15,46,47
The microbial ecology of the biofilm is another factor
influencing susceptibility. Biofilms composed of multiple spe-
cies are less susceptible than single-species biofilms, although
this is not always the case with the corresponding planktonic
cultures.19,36,37,48
Some biocides are more effective than others at inactivating
bacteria in biofilms, although conflicting data have been re-
ported, which may be explained by differences in experimental
conditions.20,24,35,49,50 In one study, susceptibility varied by
phase, organism and biocide.20 In another study, the oxidizing
agents sodium hypochlorite and peroxygens were more effec-
tive than a range of other chemicals (including alcohols,
biguanides, halogens, phenols and quaternary ammonium
compounds) for inactivating P. aeruginosa and S. aureus bio-
films.35 In other studies, sodium hypochlorite was more
Table I
Biocide susceptibility of planktonic vs surface-attached and/or biofilm mode
Author Organisms
(N isolates)
Biocides Methods Findings
Condell 201218 Salmonella
enterica (189)
Seven common food
contact surface
biocides
Tested in planktonic
culture, dried on surfaces
and as established high-
nutrient (2-day) or low-
nutrient (7-day) biofilms on
microtitre plates
Susceptibility rank: high-nutrient
biofilm < low-nutrient
biofilm < surface dried < planktonic
Behnke 201237 Pseudomonas
aeruginosa (1);
Burkholderia
cepacia (1)
Chlorine dioxide Tested in single- and
binary-species planktonic
culture, attached (4-day)
and detached biofilm
Susceptibility rank: attached
biofilm < detached
biofilm ¼ planktonic cells. Binary
cultures were less susceptible than
single-species cultures
Xing 201249 Staphylococcus
aureus (13)
Chlorhexidine and
harmaline
Tested in planktonic
culture and in 2-day
biofilms on microtitre
plates
Biofilms were 10 to >100 times less
susceptible to chlorhexidine and >2
times less susceptible to harmaline.
Synergy noted for most strains
Leung 201220 Candida spp. (4);
Escherichia coli (2)
Sodium hypochlorite,
ethanol, hydrogen
peroxide and iodine
Tested in planktonic
culture (low- and high-
titre), attached cells
(90 min) and 1-day biofilm
in microtitre plates; 24-h
and 5-min contact times
compared
Susceptibility rank:
biofilm < attached cells < high-titre
planktonic cells  low-titre
planktonic cells. MICs for biofilm vs
planktonic cells up to >10-fold
higher for 5-min and 24-h exposures
Behnke 201136 P. aeruginosa (1);
B. cepacia (1)
Sodium hypochlorite Tested in single- and
binary-species planktonic
cultures, attached (4-day)
and detached biofilms
Susceptibility rank: attached
biofilm < detached
biofilm ¼ planktonic cells. Binary-
species cultures were less
susceptible than single-species
cultures for attached and detached
biofilms, but the reverse was true for
planktonic cells
Xu 201134 Neisseria
gonorrhoeae (3)
Atmospheric
pressure non-
equilibrium plasma
Tested dried on glass
surfaces or 4-day biofilm on
glass
Bacteria in biofilm survived
approximately twice as long as
bacteria dried on surfaces
Wong 201044 S. enterica (1) Six biocides Tested in planktonic
culture or 3-day biofilm on
microtitre plates
Bacteria in biofilm were less
susceptible than planktonic cells for
all but sodium hypochlorite
Tote 201035 S. aureus (1);
P. aeruginosa (1)
12 biocides Tested in planktonic
culture or in 1-day
(P. aeruginosa) or 3-day
(S. aureus) biofilm on
microtitre plates
Most disinfectants tested did not
eliminate bacteria in the biofilm
after 60-min contact. Only hydrogen
peroxide and chlorine had an impact
on the biofilm matrix
Lee 200924 Meticillin-resistant
S. aureus (2)
Three denture-
cleaning biocides
Tested in planktonic
culture, sessile biofilm
(4 h), established biofilm
(24 h) or mature biofilm
(120 h) on resin
Two of three biocides were less
effective for the inactivation of
bacteria in biofilm. NaOCl was the
most effective against biofilm
Hendry
2009103
S. aureus (1);
meticillin-resistant
S. aureus (1);
P. aeruginosa (1);
E. coli (1); Candida
albicans (1)
Eucalyptus oil,
’1,8-cineole’ and
chlorhexidine
Tested in planktonic
culture or 2-day biofilm on
microtitre plates
Biofilm MICs and MBCs were 10 to
>100 times less susceptible than
planktonic culture. Synergy between
chlorhexidine and the other agents
was noted against some organisms
Smith 20087 Meticillin-resistant
S. aureus (8);
P. aeruginosa (8)
Benzalkonium
chloride, triclosan
and chlorhexidine
Tested in planktonic
culture or 1-day biofilms on
metal or plastic discs
MBCs for MRSA biofilms were 100 to
1000 times greater than for
planktonic cells; MBCs for
P. aeruginosa biofilm were 10 to 100
timesgreater than for planktonic cells
(continued on next page)
Table I (continued )
Author Organisms
(N isolates)
Biocides Methods Findings
Brandle 200819 Enterococcus
faecalis (1);
Streptococcus
sobrinus (1);
C. albicans (1);
Actinomyces.
naeslundii (1),
Fusobacterium
nucleatum (1)
Calcium hydroxide Tested in planktonic
culture, adherent cells,
single-species 5-day biofilm
and mixed-species 5-day
biofilm on dentin and
detached biofilm
Susceptibility rank: mixed species
biofilm < single-species biofilm <
adherent < planktonic ¼ detached
biofilm
Nett 200826 C. albicans (2);
Candida parapsilosis
(2); Candida
glabrata (1)
Ethanol, hydrogen
peroxide and sodium
dodecyl sulphate
Tested in planktonic
culture, planktonic culture
with adjustment to match
the cell density of the
biofilm and 1-day biofilm on
microtitre plates
Concentrations required to inhibit
growth in biofilm were 2- to 10-fold
higher; lower concentrations of
hydrogen peroxide prevented
biofilm formation than the other
agents tested
Karpanen
2008102
Staphylococcus
epidermidis (2)
Chlorhexidine
gluconate, tea tree
oil, eucalyptus oil
and thymol
Tested in planktonic
culture or 3-day biofilm on
microtitre plates
MICs/MBCs were elevated up to 16-
fold for biofilm; synergy was noted
between chlorhexidine and
eucalyptus oil
Bjarnsholt
2007104
P. aeruginosa (1) Silver Tested in planktonic
culture or 4-day biofilm
Biofilm was 10e100 times less
susceptible than planktonic cells
Tabak 200740 Salmonella
typhimurium (3)
Triclosan Tested in planktonic (log
and stationary phase)
culture and in 1-day biofilm
on microtitre plates
Susceptibility rank:
biofilm < stationary phase
planktonic < log phase planktonic.
8-log difference in bacteria surviving
in biofilm vs planktonic log phase
Surdeau
2006105
E. coli (1);
Enterococcus hirae
(1); P. aeruginosa
(1); S. aureus (1)
Novel disinfectant
(Oxsil 320N)
Tested in planktonic
culture and 1-day biofilm
on stainless steel
Disinfectant concentration required
to achieve a 5-log reduction was
approximately 10 times more for
biofilm vs planktonic culture
Theraud
200430
Five fungi from
patient (3) and
environment (3)
Five antiseptics,
three disinfectants
and UVC
Tested in single- and
mixed-species planktonic
culture, and single- and
mixed-species 1-day
biofilms on microtitre
plates
UVC and 3% hydrogen peroxide were
not fungicidal in initial suspension
tests. Agents were less effective
against mixed suspensions. Only
chlorhexidine was effective against
biofilms
Simoes
2003106
Pseudomonas
flourescens (1)
Ortho-
phthalaldehyde
Tested in planktonic
culture and 6-day biofilm
on glass
Biofilm was less susceptible than
planktonic cells based on respiratory
activity
Bardouniotis
200333
Mycobacterium
fortuitum (1);
Mycobacterium
marinum (1)
Seven biocides Tested in planktonic
culture and biofilm on
microtitre plate assessed
over 14 days
MBECs were up to 40-fold higher
than MBCs for M. fortuitum, but not
for M. marinum
Elvers 2002107 Alcaligenes
denitrificans (1);
Pseudomonas
alcaligenes (1);
Stenotrophomonas
maltophilia (1);
Flavobacterium
indologenes (1);
Fusarium oxysporum
(1); Fusobacterium
solani (1);
Rhodotorula
glutinis (1)
One biocide
(isothiazolone
compound)
Tested in single-species
planktonic culture, and
single- and mixed-species
1-day biofilms on glass
Biofilms were less susceptible than
planktonic cells. Mixed-species
biofilm, particularly for the bacterial
species, offered greater protection
TesTable I (continued )
Author Organisms
(N isolates)
Biocides
Peng 200231 Bacillus cereus (1) Sodium hypochloriteeffective than chlorhexidine for inactivating Enterococcus
faecium and MRSA in biofilms,24,50 whereas chlorhexidine was
found to be effective against yeast biofilms when sodium hy-
pochlorite was not effective.30 In general, oxidizing agents
target multiple biofilm components and microbial targets,
whereas other biocides such as chlorhexidine only target cell
and quaternary
ammonium
compounds
cul
sta
and
sta
wit
Bardouniotis
2001108
Mycobacterium
phlei (1)
Seven biocides Tes
cul
on
Joseph 200132 Salmonella spp. (2) Chlorine and iodine Tes
cul
on
sta
Cochran
200029
P. aeruginosa (1) Monochloramine and
hydrogen peroxide
Tes
cul
bio
and
Elasri 199953 P. aeruginosa (1) UVA, UVB and UVC Stra
cul
alg
ove
Das 199843 S. epidermidis (1);
E. coli (1)
Five biocides Tes
cul
on
Stewart
199845
Enterobacter
aerogenes (1)
Four biocides Tes
cul
den
bea
Yu 199342 Klebsiella
pneumoniae (1)
Sodium hypochlorite
and monochloramine
Tes
cul
sta
Eginton 199856 S. epidermidis (1);
P. aeruginosa (1)
Sodium hypochlorite
and dodigen; SDS and
Tween-80
Tes
cul
on
LeChevallier
198825
Pseudomonas
picketti,
Pseudomonas
paucimobilis
Moraxellaa;
K. pneumoniae (1)
Hypochlorous acid,
hypochlorite,
chlorine dioxide and
monochloramine
Tes
cul
on
car
MBC, minimum bactericidal concentration; MIC, minimum inhibitory co
ultraviolet.
Search strategy: Pubmed search for ‘susceptibility planktonic biofilm b
selected for review and 21 were included. A further 10 articles were inclu
they tested organisms and biocides relevant to disinfection in healthca
attached and/or biofilm mode susceptibility.
a Population from de-ionized water system: composition 70% P. picketMethods Findings
ted in planktonic Susceptibility rank: milk biofilmwall components; thus, oxidizing agents tend to have a higher
level of efficacy against biofilms.20,24,35,49,50 The variations in
performance of biocides under different experimental condi-
tions may have implications for practice, where the same
biocide could have a different impact on biofilms in different
settings.
ture, attached to
inless steel chips (4 h)
8-day biofilm on
inless steel with or
hout milk
< biofilm < attached < planktonic.
5-log difference between planktonic
cells and milk biofilm
ted in planktonic
ture and 5-day biofilm
microtitre plate
MBECs were higher than MBCs after
30-min and 120-min exposure to
most agents tested
ted in planktonic
ture and 10-day biofilms
plastic, cement and
inless steel
Biofilms were less susceptible to
both disinfectants; survival time no
more than 10 min in suspension
vs > 25 min in biofilm
ted in planktonic
ture and 3-h to 3-day
films on alginate beads
glass slides
Biofilms were less susceptible to
both disinfectants. Reduced
diffusion of biocide in biofilm did not
explain reduced susceptibility
in tested in planktonic
ture or biofilm in
inate beads assessed
r 1 day
Biofilm transmitted only a small
amount of UV radiation (13% of UVC,
31% of UVB and 33% of UVA),
meaning biofilm was less susceptible
than planktonic cells
ted in planktonic
ture and 6e24-h biofilms
microtitre plates
Biofilms were up to 33-fold less
susceptible to the disinfectants
tested, apart from chloroxylenol and
cetrimide (E. coli only)
ted in planktonic
ture and high- and low-
sity biofilms on alginate
ds assessed over 5 h
Susceptibility rank: high-density
biofilm < low-density
biofilm < planktonic
ted in planktonic
ture and biofilm on
inless steel discs
No difference identified between
planktonic and biofilm cells
ted in planktonic
ture and 16-h biofilms
glass and stainless steel
Biofilms were up to >1000-fold less
susceptible than planktonic cells;
attachment to the surfaces was
loosened
ted in planktonic
ture and 3-week biofilms
granular activated
bon, metal or glass
Biofilms were 150 to 3000 times less
susceptible to hypochlorous acid,
and 2- to 100-fold less susceptible to
monochloramine
ncentration; MBEC, minimal biofilm eradicating concentration; UV,
iocide’ performed on 15th November 2013. Of 44 results, 35 were
ded following review of the reference lists. Articles were included if
re facilities, and included data comparing planktonic with surface-
ti; 18% Moraxella spp.; 12% P. paucimobilis.
removal by cleaning are likely to contribute to failures in
PersistenceComparing biocides may be further confounded by the
‘doseeresponse’ type relationship that has been shown be-
tween biofilm susceptibility and biocide concentration.35,51,52
For example, one study showed that 10% hydrogen peroxide
was considerably more effective for inactivating bacteria in
biofilms compared with 6% hydrogen peroxide.51 Biofilms have
also been shown to reduce the susceptibility of microbes to
physical processes such as exposure to ultraviolet (UV) radia-
tion, most likely due to poor penetration of UV into the bio-
film.53 This may have implications for automated room
disinfection systems using UV radiation.54 To the authors’
knowledge, no studies have evaluated the impact of hydrogen-
peroxide-based automated room disinfection systems against
biofilms, although emerging data suggest that liquid hydrogen
peroxide, as an oxidizing agent, targets both the biofilm matrix
and microbes in the biofilm.35
Aside from the inactivation of microbes attached to sur-
faces, the chemical properties of biocides also seem to be
important in terms of preventing, promoting or dismantling
biofilms.26,35,55 One study showed that only sodium hypochlo-
rite and hydrogen peroxide damaged both the bacteria within
the biofilm and the biofilm matrix itself.35 Also, hydrogen
peroxide was more effective than other agents at preventing
Candida spp. biofilm formation.26 In another study, exposure to
chlorhexidine and benzalkonium chloride inhibited biofilm
formation for E. coli, K. pneumoniae and P. aeruginosa, but
promoted biofilm formation in Staphylococcus epidermidis,
suggesting that microbial factors are important.55 It is possible,
therefore, that one microbe in a biofilm may be inactivated by
a biocide, but another less susceptible microbe may survive
and then grow to replace the microbe that was inactivated.
Antibiotic susceptibility
Bacteria in biofilms are usually less susceptible to antibiotics
than bacteria in planktonic culture, and many of the mecha-
nisms for reduced susceptibility to biocides and antibiotics are
shared.5,6 Furthermore, bacteria acquired from surfaces in
biofilm mode with reduced biocide susceptibility may retain
reduced susceptibility to antibiotics.
Physical removal
The protected mode of growth offers physical protection to
cells within biofilms, and makes the physical breakdown of
biofilms challenging.5 Although biofilm attachment appears to
be loosened by some biocides,35,56 several studies have illus-
trated how difficult it can be to remove bacteria in biofilms
through cleaning and/or inactivation through disinfection. For
example, regular and extended detergent cleaning did not
remove a Bacillus cereus biofilm in vitro; a modified procedure
including heating to 70

C was required.31 Clearly, heating to
70

C is not feasible for the cleaning and disinfection of hospital
surfaces in clinical areas. Similarly, attached, viable Pseudo-
monas fragiwere detected on stainless steel surfaces after two
cleaning and disinfection procedures were tested under ‘worst-
case’ conditions at 50% in-use disinfectant concentrations.57
An acid-detergent-based method was more effective at
removing attached cells than an alkaline-detergent-based
method. However, these studies were performed using
mature biofilms which may not be representative of the bio-
films present on hospital surfaces.
Surface-attached cells and biofilms are clearly not the only
reason for failures in hospital disinfection, given the difficultyVegetative bacteria dried on to surfaces can survive for
weeks to months (or more) in vitro, despite the lack of a
nutrient source or water (aside from ambient humidity).8,9
Biofilms may explain this surprising propensity of vegetative
bacteria.8e10 This is supported by a recent study which found
that biofilm-forming strains of Acinetobacter baumannii sur-
vived for longer on dry surfaces than non-biofilm-forming
strains (36 vs 15 days; P < 0.001).64 In-vitro studies evalu-
ating the persistence of dried inocula did not supply any water
or nutrients.8e10,14,65 However, in the hospital environment,
daily and terminal cleaning or disinfection does provide a
supply of water, and some bacteria may be able to metabolize
some constituent parts of detergents and even disinfectants,
providing a nutrient source for the growth in biofilms.66e69
Transfer of plasmids and development of
antimicrobial resistance
Biofilms are suited for horizontal gene dissemination
because they are a mixed population at high bacterial density,
which facilitates metabolic activity in the harshest environ-
ments, albeit at a reduced rate. Horizontal transfer of plasmids
does occur through conjugation, as illustrated by the transfer
of extended-spectrum b-lactamase (CTX-M-15) and carbape-
nemase (NDM-1) plasmids between Enterobacteriaceae when
dried on surfaces.70,71 Furthermore, the mutation rate (the
rate at which DNA replication mistakes occur during cell divi-
sion) of bacteria in biofilms is increased.6,72 Thus, both hori-
zontal transfer of resistance determinants such as plasmids and
increased mutation rates could result in the acquisition or de-
novo development of reduced susceptibility to antimicrobial
agents and other important microbial capabilities, such as
increased virulence.
Tackling surface-attached cells and biofilms
Surface-attached cells, especially established biofilms, pre-hospital cleaning. This could partly explain why disinfectants
that are effective for the inactivation of planktonic bacteria in
laboratory tests are not effective for the eradication of a
considerably lower load of the same bacterial species from
hospital surfaces.11,13,58e60 In support of this, it is noteworthy
that the biofilms identified by Vickery et al. were on surfaces
that had been cleaned with detergent and then disinfected
using 500 ppm chlorine.15 These findings may have implications
for infection control practices within hospitals, and on the
choice of appropriate disinfectants used to decontaminate
surfaces.11,13,54
The presence of biofilms on dry hospital surfaces could also
interfere with attempts to recover microbes through environ-
mental sampling.15,61e63 This could mean that an environ-
mental reservoir of a pathogen remains undetected, or the
concentration of contamination and degree of associated risk is
underestimated.in achieving adequate distribution and contact time using
manual methods.11,13,54,58 However, both reduced biocide
susceptibility (Table I) and increasing resilience to physicalsent a difficult challenge to hospital cleaning and disinfection,
combining protection from physical removal with reduced sus-
ceptibility to biocides (Table I).31,57 A number of different ap-
proaches are available to tackle surface-attached cells and
biofilms. Using physical methods to dislodge detached bacteria,
which can be aided by the use of a detergent, can be effective in
removing established biofilms and preventing the development
of biofilms.5,56,73 However, detergent cleaning alonemay not be
sufficient to remove biofilms.5,15,31,56,61,73 Tackling themicrobes
in the biofilm alone (e.g. using some disinfectants or attempts to
interfere with quorum sensing) can be effective, but may not
reach microbes protected deep in the biofilm matrix. Tackling
the biofilm matrix alone (e.g. using enzymatic digestion) will
help to reach microbes protected within the biofilm matrix and
interrupt persistenceof thebiofilm, butwill not necessarily have
direct microbicidal activity. Thus, tackling both the microbes in
the biofilm and the biofilm matrix simultaneously (using
oxidizing disinfectants or combination approaches) offers the
potential to reach microbes protected deep in the matrix and
interrupt the persistence of the biofilm. In addition, some bio-
cides have the ability to reduce biofilm formation, which can be
assisted by choosing surface materials that do not readily sup-
port biofilm formation.
Biocides and biocide adjuvants
Differences between biocides appear to influence their ac-
tivity against bacteria attached to surfaces and may also pro-
mote, prevent or dismantle biofilms. Thus, biocides with the
highest activity against bacteria attached to surfaces, and
ideally those with the ability to prevent biofilm formation and
dismantle existing biofilms, should be selected. Emerging data
indicate that oxidizing agents may possess more of these
properties than other agents.35 Similarly, detergent formula-
tions that are better at physical removal should be selected,
although there is a paucity of data on the capacity of currently
available detergents to address surface-attached cells.73
Several novel approaches also warrant consideration as po-
tential additives to hospital detergents or disinfectants to
augment their effectiveness against biofilms. Firstly, certain
enzymes such as DNase and dispersinB have been shown to
dissolve the biofilm matrix.73e78 For example, detergents sup-
plemented with high concentrations of enzymes were effective
against hydrated biofilms, whereas detergents supplemented
with low concentrations of enzymes were not.73 Secondly,
quorum-sensing inhibitors have proven successful in increasing
antimicrobial susceptibility.79e81 In one study, drimendiol, a
quorum-sensing inhibitor, was found to enhance the effects of
copper sulphate on biofilms of Pseudomonas syringae.81 Thirdly,
recently discovered human antimicrobial peptides also have
antibiofilm activities.82e84 For example, a range of antimicro-
bial peptides tested against multi-drug-resistant A. baumannii
demonstrated direct antimicrobial activity, and enhanced the
activity of a range of other antimicrobial agents.82
However, the addition of enzymes, quorum-sensing in-
hibitors or antimicrobial peptides into a cleaning or disinfection
solution would result in chemical residues on surfaces with
associated health and safety implications, so are not recom-
mended without further study. Another approach is the inclu-
sion of bacteriophages, which have been found to disrupt
biofilms.85,86 For example, Streptococcus pyogenes biofilms
were degraded by PlyC, a bacteriophage-encoded endolysin,
which also acted synergistically with a range of antimicrobialagents.86 However, the therapeutic use of bacteriophages in
human medicine and, by implication, in the clinical environ-
ment is controversial due topotential for the rapid development
of resistance and the risk that the introduced bacteriophages
may play an unintended role in horizontal gene transfer.85,87
Surface modification to prevent biofilm formation
Some surface materials are more prone to biofilm formation
than others.71,88 A recent study reviewed attempts to modify
the chemical or physical surface properties of medical devices
to inhibit or prevent microbial adhesion.88 These include
‘liquid glass’ (silicon dioxide), Sharklet pattern,89,90 advanced
polymer coatings [e.g. polyethylene glycol (PEG), super-
hydrophobic/philic and zwitterionic]9194 and diamond-like
carbon films.95 Whilst these technologies have the potential
to reduce biofilm deposition on hospital surfaces, they are at
an early stage of development. The feasibility and cost-
effectiveness of scaling up these technologies for use on hos-
pital surfaces needs to be evaluated.
Another approach is the implementation of antimicrobial
surfaces. Options include metals such as copper and silver, or
chemicals such as organosilanes with quaternary ammonium
groups and light-activated antimicrobials.12,71,96 Copper is the
most-studied candidate for antimicrobial surfaces, and has
been shown to inactivate microbes and DNA deposited on sur-
faces and may reduce the transmission of pathogens in the
hospital setting.12,71,97 However, the presence of a condition-
ing film can greatly reduce the efficacy of antimicrobial
surfaces.98e100 Thus, an antimicrobial surface that combines
reduced biofilm formation with direct antimicrobial activity is
a promising area for future research. Another challenge in
developing an antimicrobial surface for hospitals is the
requirement for multiple different surface types (from fabric
to hard surfaces) with a range of required functions. Thus,
there is unlikely to be a single agent or surface structure that is
suitable for all applications.
Implications for susceptibility testing
Surface-attached cells and biofilms are a more accurate
reflection of the occurrence of bacteria in nature than plank-
tonic cells.1,101 However, planktonic culture remains the cur-
rent model for many microbiological studies and testing
standards including susceptibility testing.1,101 Although quan-
titative surface tests for evaluation of the bactericidal activity
of chemical disinfectants do exist (e.g. BS EN 13697:2001),
none have been published for EPS-producing biofilms. Future
testing should specify the use of surface-attached cells and
consider the use of biofilm models to ensure that the disin-
fectants tested are as effective in the ‘real world’ as in labo-
ratory tests.1 It seems likely that low-nutrient, low-density
surface-attached cells would be more appropriate than high-
nutrient, high-density established biofilms. Most in-vitro
studies measured growth over a 24-h period to evaluate the
impact of a chemical biocide to determine the MIC or MBC,
using methodology often used to test antibiotic susceptibility
(Table I).7,20,102 One study compared the MICs of four common
biocides for E. coil and various Candida spp. with a ‘contact
time’ of 5 min and 24 h.20 Unsurprisingly, the concentration
required to inhibit growth within 5 min was considerably
greater than the concentration required to inhibit growth over
19. Brandle N, Zehnder M, Weiger R, Waltimo T. Impact of growth
conditions on susceptibility of five microbial species to alkaline24 h. Thus, as biocides are only applied for a short period in
practice, evaluating the impact of a biocide over a short con-
tact time as per most published biocide testing standards is
more suitable for in-vitro biocide studies than measuring the
MIC or MBC when microbes are grown in varying concentrations
of biocide.
Further research is required to evaluate the prevalence and
composition of biofilms in situ on hard and soft hospital sur-
faces, to develop in-vitro models that are representative of
those likely to be found on hospital surfaces, and to optimize
methods to tackle biofilms on hospital surfaces, which may
include new cleaning and disinfection agents and adjuvants,
new technologies (such as microfibre or automated room
disinfection technology), and surface modification.15
Conclusion
Surface-attached cells are likely to be common on dry hos-
pital surfaces, and there is evidence that they also harbour
established biofilms. The variety of methods used to create and
evaluate in-vitro biofilms makes it difficult to compare studies
evaluating antibiofilm biocide activity. Nonetheless, microbes
attached to surfaces, especially established biofilms, are less
susceptible to chemical biocides, UV radiation and antibiotics
than their corresponding planktonic bacteria. The phase of the
surface-attached microbes influences susceptibility: attached
cells are more susceptible to biocides than established bio-
films; low-density, nutrient-limited biofilms make less of an
impact on biocide susceptibility than high-density, high-
nutrient biofilms; and biocides are less effective for inacti-
vating bacteria in mixed-species biofilms than in single-species
biofilms. Biocide-specific issues also influence susceptibility in
terms of activity against bacteria in biofilms, and the preven-
tion, promotion and dismantling of biofilms. Reduced suscep-
tibility to biocides combined with protection from physical
removal through cleaning is likely to contribute to failures in
hospital cleaning and disinfection.
Biofilms may explain why vegetative bacteria can survive for
unusually long periods (weeks to months) on dry hospital sur-
faces. Also, the presence of surface-attached bacteria and bio-
films is likely to interferewith attempts to recover bacteria from
hospital surfaces, and may lead to underestimation of both the
prevalence of contamination with pathogens and the number of
bacteria that are on surfaces. This has important implications,
particularly for hospital outbreak investigation. Biofilms provide
a mixed bacterial community where the horizontal transfer of
resistance genes may occur. Attempts to tackle surface-
attached microbes and biofilms on hospital surfaces should
include: identification and selection of biocide and detergents
with the best all-round performance, including the ability to
inactivate surface-attached cells and biofilms; ensuring that in-
vitro tests are developed to model surface-attached microbes
likely to be encountered in the field; harnessing surface science
to develop a hospital environment that reduces the chance of
biofilm formation; and further research to develop novel ap-
proaches to augment the activity of biocides against surface-
attached microbes, including established biofilms.
Conflict of interest statement
JAO is employed part-time by Bioquell, and JAGS, JC and SY
are employed by Bioquell. All other authors have no con-
flicts of interest to declare.stress. J Endod 2008;34:579e582.
20. Leung CY, Chan YC, Samaranayake LP, Seneviratne CJ. BiocideFunding source
None.
References
1. Lindsay D, von Holy A. Bacterial biofilms within the clinical
setting: what healthcare professionals should know. J Hosp Infect
2006;64:313e325.
2. Hall-Stoodley L, Stoodley P. Biofilm formation and dispersal and the
transmission of human pathogens. Trends Microbiol 2005;13:7e10.
3. Hall-Stoodley L, Costerton JW, Stoodley P. Bacterial biofilms:
from the natural environment to infectious diseases. Nat Rev
Microbiol 2004;2:95e108.
4. Loveday HP, Wilson J, Kerr K, Pitchers R, Walker JT, Browne J.
Pseudomonas infection and healthcare water systems e a rapid
systematic review. J Hosp Infect 2014;86:7e15.
5. Mah TF, O’Toole GA. Mechanisms of biofilm resistance to anti-
microbial agents. Trends Microbiol 2001;9:34e39.
6. Hoiby N, Bjarnsholt T, Givskov M, Molin S, Ciofu O. Antibiotic
resistance of bacterial biofilms. Int J Antimicrob Agents
2010;35:322e332.
7. Smith K, Hunter IS. Efficacy of common hospital biocides with
biofilms of multi-drug resistant clinical isolates. J Med Microbiol
2008;57:966e973.
8. Otter JA, French GL. Survival of nosocomial bacteria and spores
on surfaces and inactivation by hydrogen peroxide vapor. J Clin
Microbiol 2009;47:205e207.
9. Kramer A, Schwebke I, Kampf G. How long do nosocomial path-
ogens persist on inanimate surfaces? A systematic review. BMC
Infect Dis 2006;6:130.
10. Wagenvoort JH, De Brauwer EI, Penders RJ, Willems RJ, Top J,
Bonten MJ. Environmental survival of vancomycin-resistant
Enterococcus faecium. J Hosp Infect 2011;77:282e283.
11. Otter JA, Yezli S, French GL. The role played by contaminated
surfaces in the transmission of nosocomial pathogens. Infect
Control Hosp Epidemiol 2011;32:687e699.
12. Salgado CD, Sepkowitz KA, John JF, et al. Copper surfaces reduce
the rate of healthcare-acquired infections in the intensive care
unit. Infect Control Hosp Epidemiol 2013;34:479e486.
13. Donskey CJ. Does improving surface cleaning and disinfection
reduce health care-associated infections? Am J Infect Control
2013;41:S12eS19.
14. Wagenvoort JH, Joosten EJ. An outbreak Acinetobacter bau-
mannii that mimics MRSA in its environmental longevity. J Hosp
Infect 2002;52:226e227.
15. Vickery K, Deva A, Jacombs A, Allan J, Valente P, Gosbell IB.
Presence of biofilm containing viable multiresistant organisms
despite terminal cleaning on clinical surfaces in an intensive care
unit. J Hosp Infect 2012;80:52e55.
16. Yezli S, Otter JA. Does the discovery of biofilms on dry hospital
environmental surfaces change the way we think about hospital
disinfection? J Hosp Infect 2012;81:293e294.
17. Smith K, Perez A, Ramage G, Gemmell CG, Lang S. Comparison of
biofilm-associated cell survival following in vitro exposure of
meticillin-resistant Staphylococcus aureus biofilms to the anti-
biotics clindamycin, daptomycin, linezolid, tigecycline and van-
comycin. Int J Antimicrob Agents 2009;33:374e378.
18. Condell O, IversenC,Cooney S, et al. Efficacy of biocides used in the
modern food industry to control Salmonella enterica, and links be-
tween biocide tolerance and resistance to clinically relevant anti-
microbial compounds. Appl Environ Microbiol 2012;78:3087e3097.resistance of Candida and Escherichia coli biofilms is associated
with higher antioxidative capacities. JHosp Infect 2012;81:79e86.
21. Parsek MR, Greenberg EP. Sociomicrobiology: the connections
between quorum sensing and biofilms. Trends Microbiol
2005;13:27e33.
22. Stewart PS. Diffusion in biofilms. J Bacteriol 2003;185:1485e1491.
23. Keren I, Kaldalu N, Spoering A, Wang Y, Lewis K. Persister cells
and tolerance to antimicrobials. FEMS Microbiol Lett
2004;230:13e18.
24. Lee D, Howlett J, Pratten J, et al. Susceptibility of MRSA biofilms
to denture-cleansing agents. FEMS Microbiol Lett
2009;291:241e246.
25. LeChevallier MW, Cawthon CD, Lee RG. Inactivation of biofilm
bacteria. Appl Environ Microbiol 1988;54:2492e2499.
26. Nett JE, Guite KM, Ringeisen A, Holoyda KA, Andes DR. Reduced
biocide susceptibility in Candida albicans biofilms. Antimicrob
Agents Chemother 2008;52:3411e3413.
27. Watamoto T, Samaranayake LP, Egusa H, Yatani H,
Samaranayke YH, Seneviratne CJ. Susceptibility of Candida
albicans filamentation-defective mutants to clinical biocides.
J Hosp Infect 2010;74:189e191.
28. Seneviratne CJ, Jin LJ, Samaranayake YH, Samaranayake LP. Cell
density and cell aging as factors modulating antifungal resistance
of Candida albicans biofilms. Antimicrob Agents Chemother
2008;52:3259e3266.
29. Cochran WL, McFeters GA, Stewart PS. Reduced susceptibility of
thin Pseudomonas aeruginosa biofilms to hydrogen peroxide and
monochloramine. J Appl Microbiol 2000;88:22e30.
30. Theraud M, Bedouin Y, Guiguen C, Gangneux JP. Efficacy of an-
tiseptics and disinfectants on clinical and environmental yeast
isolates in planktonic and biofilm conditions. J Med Microbiol
2004;53:1013e1018.
31. Peng JS, Tsai WC, Chou CC. Inactivation and removal of Bacillus
cereus by sanitizer and detergent. Int J Food Microbiol
2002;77:11e18.
32. Joseph B, Otta SK, Karunasagar I, Karunasagar I. Biofilm forma-
tion by Salmonella spp. on food contact surfaces and their
sensitivity to sanitizers. Int J Food Microbiol 2001;64:367e372.
33. Bardouniotis E, Ceri H, Olson ME. Biofilm formation and biocide
susceptibility testing of Mycobacterium fortuitum and Mycobac-
terium marinum. Curr Microbiol 2003;46:28e32.
34. Xu L, Tu Y, Yu Y, Tan M, Li J, Chen H. Augmented survival of
Neisseria gonorrhoeae within biofilms: exposure to atmospheric
pressure non-thermal plasmas. Eur J Clin Microbiol Infect Dis
2011;30:25e31.
35. Tote K, Horemans T, Vanden Berghe D, Maes L, Cos P. Inhibitory
effect of biocides on the viable masses and matrices of Staphy-
lococcus aureus and Pseudomonas aeruginosa biofilms. Appl En-
viron Microbiol 2010;76:3135e3142.
36. Behnke S, Parker AE, Woodall D, Camper AK. Comparing the
chlorine disinfection of detached biofilm clusters with those of
sessile biofilms and planktonic cells in single- and dual-species
cultures. Appl Environ Microbiol 2011;77:7176e7184.
37. Behnke S, Camper AK. Chlorine dioxide disinfection of single and
dual species biofilms, detached biofilm and planktonic cells.
Biofouling 2012;28:635e647.
38. Bjergbaek LA, Haagensen JA, Molin S, Roslev P. Effect of oxygen
limitation and starvation on the benzalkonium chloride suscepti-
bility of Escherichia coli. J Appl Microbiol 2008;105:1310e1317.
39. Field TR, White A, Elborn JS, Tunney MM. Effect of oxygen limi-
tation on the in vitro antimicrobial susceptibility of clinical iso-
lates of Pseudomonas aeruginosa grown planktonically and as
biofilms. Eur J Clin Microbiol Infect Dis 2005;24:677e687.
40. Tabak M, Scher K, Hartog E, et al. Effect of triclosan on Salmo-
nella typhimurium at different growth stages and in biofilms.
FEMS Microbiol Lett 2007;267:200e206.
41. Smith K, Gemmell CG, Hunter IS. The association between
biocide tolerance and the presence or absence of qac genes
among hospital-acquired and community-acquired MRSA isolates.
J Antimicrob Chemother 2008;61:78e84.42. Yu FP, Pyle BH, McFeters GA. A direct viable count method for
the enumeration of attached bacteria and assessment of biofilm
disinfection. J Microbiol Methods 1993;17:167e180.
43. Das JR, Bhakoo M, Jones MV, Gilbert P. Changes in the biocide
susceptibility of Staphylococcus epidermidis and Escherichia coli
cells associated with rapid attachment to plastic surfaces. J Appl
Microbiol 1998;84:852e858.
44. Wong HS, Townsend KM, Fenwick SG, Trengove RD,
O’Handley RM. Comparative susceptibility of planktonic and 3-
day-old Salmonella typhimurium biofilms to disinfectants.
J Appl Microbiol 2010;108:2222e2228.
45. Stewart PS, Grab L, Diemer JA. Analysis of biocide transport
limitation in an artificial biofilm system. J Appl Microbiol
1998;85:495e500.
46. van de Mortel M, Halverson LJ. Cell envelope components
contributing to biofilm growth and survival of Pseudomonas putida
in low-water-content habitats. Mol Microbiol 2004;52:735e750.
47. Flemming HC, Wingender J. The biofilm matrix. Nat Rev Micro-
biol 2010;8:623e633.
48. Bridier A, Sanchez-Vizuete Mdel P, Le Coq D, et al. Biofilms of a
Bacillus subtilis hospital isolate protect Staphylococcus aureus
from biocide action. PLoS One 2012;7:e44506.
49. Xing M, Shen F, Liu L, et al. Antimicrobial efficacy of the alkaloid
harmaline alone and in combinationwith chlorhexidine digluconate
against clinical isolates of Staphylococcus aureus grown in plank-
tonic and biofilm cultures. Lett Appl Microbiol 2012;54:475e482.
50. Williamson AE, Cardon JW, Drake DR. Antimicrobial susceptibility
of monoculture biofilms of a clinical isolate of Enterococcus
faecalis. J Endod 2009;35:95e97.
51. Yun HS, Kim Y, Oh S, Jeon WM, Frank JF, Kim SH. Susceptibility of
Listeria monocytogenes biofilms and planktonic cultures to
hydrogen peroxide in food processing environments. Biosci Bio-
technol Biochem 2012;76:2008e2013.
52. Peeters E, Nelis HJ, Coenye T. Evaluation of the efficacy of
disinfection procedures against Burkholderia cenocepacia bio-
films. J Hosp Infect 2008;70:361e368.
53. Elasri MO, Miller RV. Study of the response of a biofilm bacterial
community to UV radiation. Appl Environ Microbiol
1999;65:2025e2031.
54. Otter JA, Yezli S, Perl TM, Barbut F, French GL. Is there a role for
‘no-touch’ automated room disinfection systems in infection
prevention and control? J Hosp Infect 2013;83:1e13.
55. Houari A, Di Martino P. Effect of chlorhexidine and benzalkonium
chloride on bacterial biofilm formation. Lett Appl Microbiol
2007;45:652e656.
56. Eginton PJ, Holah J, Allison DG, Handley PS, Gilbert P. Changes in
the strength of attachment of micro-organisms to surfaces
following treatment with disinfectants and cleansing agents. Lett
Appl Microbiol 1998;27:101e105.
57. Stone LS, Zottola EA. Effect of cleaning and sanitizing on the
attachment of Pseudomonas fragi to stainless steel. J Food Sci
1985;50:951e956.
58. Manian FA, Griesenauer S, Senkel D, et al. Isolation of Acineto-
bacter baumannii complex and methicillin-resistant Staphylo-
coccus aureus from hospital rooms following terminal cleaning
and disinfection: can we do better? Infect Control Hosp Epi-
demiol 2011;32:667e672.
59. Goodman ER, Platt R, Bass R, Onderdonk AB, Yokoe DS, Huang SS.
Impact of an environmental cleaning intervention on the pres-
ence of methicillin-resistant Staphylococcus aureus and
vancomycin-resistant enterococci on surfaces in intensive care
unit rooms. Infect Control Hosp Epidemiol 2008;29:593e599.
60. Wilcox MH, Fawley WN, Wigglesworth N, Parnell P, Verity P,
Freeman J. Comparison of the effect of detergent versus hypo-
chlorite cleaning on environmental contamination and incidence
of Clostridium difficile infection. J Hosp Infect 2003;54:109e114.
61. French GL, Otter JA, Shannon KP, Adams NM, Watling D,
Parks MJ. Tackling contamination of the hospital environment by
methicillin-resistant Staphylococcus aureus (MRSA): a comparison
between conventional terminal cleaning and hydrogen peroxide
vapour decontamination. J Hosp Infect 2004;57:31e37.
62. Zanetti G, Blanc DS, Federli I, et al. Importation of Acinetobacter
baumannii into a burn unit: a recurrent outbreak of infection
associated with widespread environmental contamination. Infect
Control Hosp Epidemiol 2007;28:723e725.
63. Boyce JM, Havill NL, Havill HL, Mangione E, Dumigan DG,
Moore BA. Comparison of fluorescent marker systems with 2
quantitative methods of assessing terminal cleaning practices.
Infect Control Hosp Epidemiol 2011;32:1187e1193.
64. Espinal P, Marti S, Vila J. Effect of biofilm formation on the
survival of Acinetobacter baumannii on dry surfaces. J Hosp
Infect 2012;80:56e60.
65. Wagenvoort JH, Sluijsmans W, Penders RJ. Better environmental
survival of outbreak vs. sporadic MRSA isolates. J Hosp Infect
2000;45:231e234.
66. Weber DJ, Rutala WA, Sickbert-Bennett EE. Outbreaks associated
with contaminated antiseptics and disinfectants. Antimicrob
Agents Chemother 2007;51:4217e4224.
67. BasuA,Dixit SS, Phale PS.Metabolismof benzyl alcohol via catechol
ortho-pathway in methylnaphthalene-degrading Pseudomonas
putida CSV86. Appl Microbiol Biotechnol 2003;62:579e585.
68. Gottsching A, Schmidt S. Productive degradation of the biocide
benzylbenzoate by Acinetobacter sp. strain AG1 isolated from
the River Elbe. Res Microbiol 2007;158:251e257.
69. Ying GG. Fate, behavior and effects of surfactants and their degra-
dation products in the environment. Environ Int 2006;32:417e431.
70. Molin S, Tolker-Nielsen T. Gene transfer occurs with enhanced
efficiency in biofilms and induces enhanced stabilisation of the
biofilm structure. Curr Opin Biotechnol 2003;14:255e261.
71. Warnes SL, Highmore CJ, Keevil CW. Horizontal transfer of
antibiotic resistance genes on abiotic touch surfaces: implica-
tions for public health. MBio 2012;3:e00489ee00512.
72. Driffield K, Miller K, Bostock JM, O’Neill AJ, Chopra I. Increased
mutability of Pseudomonas aeruginosa in biofilms. J Antimicrob
Chemother 2008;61:1053e1056.
73. Vickery K, Pajkos A, Cossart Y. Removal of biofilm from endo-
scopes: evaluation of detergent efficiency. Am J Infect Control
2004;32:170e176.
74. Brindle ER, Miller DA, Stewart PS. Hydrodynamic deformation and
removal of Staphylococcus epidermidis biofilms treated with
urea, chlorhexidine, iron chloride, or DispersinB. Biotechnol
Bioeng 2011;108:2968e2977.
75. Kaplan JB, LoVetri K, Cardona ST, et al. Recombinant human
DNase I decreases biofilm and increases antimicrobial suscepti-
bility in staphylococci. J Antibiot (Tokyo) 2012;65:73e77.
76. Li W, Liu H, Xu Q. Extracellular dextran and DNA affect the for-
mation of Enterococcus faecalis biofilms and their susceptibility
to 2% chlorhexidine. J Endod 2012;38:894e898.
77. Latimer J, Forbes S, McBain AJ. Attenuated virulence and bio-
film formation in Staphylococcus aureus following sublethal
exposure to triclosan. Antimicrob Agents Chemother
2012;56:3092e3100.
78. Alipour M, Suntres ZE, Omri A. Importance of DNase and alginate
lyase for enhancing free and liposome encapsulated amino-
glycoside activity against Pseudomonas aeruginosa. J Antimicrob
Chemother 2009;64:317e325.
79. Liaqat I, Bachmann RT, Sabri AN, Edyvean RG, Biggs CA. Inves-
tigating the effect of patulin, penicillic acid and EDTA on biofilm
formation of isolates from dental unit water lines. Appl Microbiol
Biotechnol 2008;81:349e358.
80. Bjarnsholt T, Jensen PO, Rasmussen TB, et al. Garlic blocks quorum
sensing and promotes rapid clearing of pulmonary Pseudomonas
aeruginosa infections. Microbiology 2005;151:3873e3880.
81. Paza C, Carcamo G, Silva M, Becerra J, Urrutia H, Sossa K. Dri-
mendiol, a drimane sesquiterpene with quorum sensing inhibition
activity. Nat Prod Commun 2013;8:147e148.82. Gopal R, Kim YG, Lee JH, et al. Synergistic effects and antibiofilm
properties of chimeric peptides against multidrug-resistant Aci-
netobacter baumannii strains. Antimicrob Agents Chemother
2014;58:1622e1629.
83. Feng X, Sambanthamoorthy K, Palys T, Paranavitana C. The hu-
man antimicrobial peptide LL-37 and its fragments possess both
antimicrobial and antibiofilm activities against multidrug-
resistant Acinetobacter baumannii. Peptides 2013;49:131e137.
84. Forbes S, McBain AJ, Felton-Smith S, Jowitt TA, Birchenough HL,
Dobson CB. Comparative surface antimicrobial properties of
synthetic biocides and novel human apolipoprotein E derived
antimicrobial peptides. Biomaterials 2013;34:5453e5464.
85. Sillankorva S, Azeredo J. Bacteriophage attack as an anti-biofilm
strategy. Methods Mol Biol 2014;1147:277e285.
86. Shen Y, Koller T, Kreikemeyer B, Nelson DC. Rapid degradation of
Streptococcus pyogenes biofilms by PlyC, a bacteriophage-
encoded endolysin. J Antimicrob Chemother 2013;68:1818e1824.
87. Lu TK, Koeris MS. The next generation of bacteriophage therapy.
Curr Opin Microbiol 2011;14:524e531.
88. Desrousseaux C, Sautou V, Descamps S, Traore O. Modification of
the surfaces of medical devices to prevent microbial adhesion
and biofilm formation. J Hosp Infect 2013;85:87e93.
89. Chung KK, Schumacher JF, Sampson EM, Burne RA, Antonelli PJ,
Brennan AB. Impact of engineered surface microtopography on
biofilm formation of Staphylococcus aureus. Biointerphases
2007;2:89e94.
90. Reddy ST, Chung KK, McDaniel CJ, Darouiche RO, Landman J,
Brennan AB. Micropatterned surfaces for reducing the risk of
catheter-associated urinary tract infection: an in vitro study on
the effect of Sharklet micropatterned surfaces to inhibit bacte-
rial colonization and migration of uropathogenic Escherichia coli.
J Endourol 2011;25:1547e1552.
91. Park KD, Kim YS, Han DK, et al. Bacterial adhesion on PEG
modified polyurethane surfaces. Biomaterials 1998;19:851e859.
92. Hou S, Burton EA, Simon KA, Blodgett D, Luk YY, Ren D. Inhibition
of Escherichia coli biofilm formation by self-assembled mono-
layers of functional alkanethiols on gold. Appl Environ Microbiol
2007;73:4300e4307.
93. Hirota K, Murakami K, Nemoto K, Miyake Y. Coating of a surface
with 2-methacryloyloxyethyl phosphorylcholine (MPC) co-
polymer significantly reduces retention of human pathogenic
microorganisms. FEMS Microbiol Lett 2005;248:37e45.
94. Cheng G, Zhang Z, Chen S, Bryers JD, Jiang S. Inhibition of bac-
terial adhesion and biofilm formation on zwitterionic surfaces.
Biomaterials 2007;28:4192e4199.
95. Liu C, Zhao Q, Liu Y, Wang S, Abel EW. Reduction of bacterial
adhesion on modified DLC coatings. Colloids Surf B Biointerfaces
2008;61:182e187.
96. Humphreys H. Self-disinfecting and microbiocide-impregnated
surfaces and fabrics: what potential in interrupting the spread of
healthcare-associated infection? Clin Infect Dis 2014;58:848e853.
97. Grass G, Rensing C, Solioz M. Metallic copper as an antimicrobial
surface. Appl Environ Microbiol 2011;77:1541e1547.
98. Ojeil M, Jermann C, Holah J, Denyer SP, Maillard JY. Evaluation of
new in vitro efficacy test for antimicrobial surface activity
reflecting UK hospital conditions. J Hosp Infect 2013;85:274e281.
99. O’Gorman J, Humphreys H. Application of copper to prevent and
control infection. Where are we now? J Hosp Infect
2012;81:217e223.
100. Airey P, Verran J. Potential use of copper as a hygienic surface;
problems associated with cumulative soiling and cleaning. J Hosp
Infect 2007;67:271e277.
101. Parsek MR, Fuqua C. Biofilms 2003: emerging themes and chal-
lenges in studies of surface-associated microbial life. J Bacteriol
2004;186:4427e4440.
102. Karpanen TJ, Worthington T, Hendry ER, Conway BR,
Lambert PA. Antimicrobial efficacy of chlorhexidine digluconate
alone and in combination with eucalyptus oil, tea tree oil and
thymol against planktonic and biofilm cultures of Staphylococcus
epidermidis. J Antimicrob Chemother 2008;62:1031e1036.
103. Hendry ER, Worthington T, Conway BR, Lambert PA. Antimicro-
bial efficacy of eucalyptus oil and 1,8-cineole alone and in
combination with chlorhexidine digluconate against microor-
ganisms grown in planktonic and biofilm cultures. J Antimicrob
Chemother 2009;64:1219e1225.
104. Bjarnsholt T, Kirketerp-Moller K, Kristiansen S, et al. Silver against
Pseudomonas aeruginosa biofilms. APMIS 2007;115:921e928.
105. Surdeau N, Laurent-Maquin D, Bouthors S, Gelle MP. Sensitivity of
bacterial biofilms and planktonic cells to a new antimicrobial
agent, Oxsil 320N. J Hosp Infect 2006;62:487e493.
106. Simoes M, Carvalho H, Pereira MO, Vieira MJ. Studies on the
behaviour of Pseudomonas fluorescens biofilms after ortho-
phthalaldehyde treatment. Biofouling 2003;19:151e157.
107. Elvers KT, Leeming K, Lappin-Scott HM. Binary and mixed popu-
lation biofilms: time-lapse image analysis and disinfection with
biocides. J Ind Microbiol Biotechnol 2002;29:331e338.
108. Bardouniotis E, Huddleston W, Ceri H, Olson ME. Characterization
of biofilm growth and biocide susceptibility testing of Mycobac-
terium phlei using the MBEC assay system. FEMS Microbiol Lett
2001;203:263e267.