Download review manuscript PY - Final Version Nov 2014

Implications and current control strategies for ventilator-associated pneumonia
Ching-Yee Loo1.2, Wing-Hin Lee1,2, Paul M. Young2,3, Rosalia Cavaliere4, Cynthia B.
Whitchurch4, and Ramin Rohanizadeh1*
1
Advanced Drug Delivery Group, Faculty of Pharmacy, University of Sydney, Sydney,
NSW 2006, Australia
2
Woolcook Institute of Medical Research and Discipline of Pharmacology, Sydney
Medical School, University of Sydney, NSW 2006, Australia
3
Discipline of Pharmacology, Sydney Medical School, University of Sydney, NSW 2006,
Australia
4
The ithree institute, University of Technology Sydney, Ultimo, NSW 2007, Australia.
*Corresponding author:
Dr Ramin Rohanizadeh
Faculty of Pharmacy (A15), University of Sydney, Sydney, New South Wales 2006,
Australia. E-mail: [email protected]
Summary
Ventilator-associated pneumonia (VAP) remains a major burden to the healthcare
system and intubated patients in intensive care units (ICU). In fact, VAP is responsible for
at least 50% of prescribed antibiotics to patients who need mechanical ventilation. One of
the factors contributing to VAP pathogenesis is believed to be rapid colonization of
biofilm-forming pathogens such as Pseudomonas aeruginosa and Staphylococcus aureus
on the surface of inserted endotracheal tubes. These biofilms serve as a protective
environment for bacterial colonies and provide enhanced resistance towards many
antibiotics. Several strategies have been adopted to reduce the occurrence of VAP such as
control of cuff pressure, aspiration of subglotic secretion and use of endotracheal tubes
with ultrathin cuff membranes, to eliminate or prevent biofilm formation. This review
presents and discusses an overview of current strategies to inhibit the colonization and
formation of biofilm on endotracheal tubes, including antibiotic treatment, surface
modification and antimicrobial agent incorporation onto endotracheal tube materials.
Keywords: biofilm, Pseudomonas aeruginosa, Staphylococcus aureus, ventilator
associated pneumonia, silver nanoparticle, surface modification
1. Introduction
Hospital-acquired pneumonia (HAP) is the second most common lung infection
caused by microorganisms and pathogens, and is responsible for one quarter of infections
in intensive care units (ICU) [1,2]. Ventilator-associated pneumonia (VAP) is defined as a
nosocomial infection occurring in patients that rely on mechanical ventilators, via invasive
methods (tracheostomy tube and endotracheal intubation). VAP accounts for more than
80% of HAP [2]. Statistics have shown that VAP affects up to 28% of intubated patients
and the incidence rate escalates with time [1,3]. The probability of developing VAP is the
highest during early intubation; with ~3% infection/day for the first 5 days of intubation
decreasing to 2% per day from day 5 to 10 [1,4].
Numerous studies have highlighted the role of the endotracheal tubes in the
pathogenesis of VAP [5,6]. In the case of critically ill patients, endotracheal tubes are used
to provide ventilation to patients. However, these endotracheal tubes often impair normal
mucocilliary clearance, leading to accumulation of tracheobronchial secretions whilst
increasing the risk of pneumonia infection [7]. Secondly, the action of endotracheal tube
insertion may cause injury and introduce exogenous and/or endogenous bacterial flora into
the mucosa [8]. This provides an active reservoir for bacterial colonization, which can
develop into multiple-antibiotic resistant biofilms [9]. A common hypothesis is that the
biofilm arise from aspirated secretions, environmental contamination of “breathe-air”
through ventilators and accumulation of secretions into tracheobronchial sections [9-11].
Irrespective of the contamination source, it is agreed that the biofilm harbors pathogenic
microorganisms that cause systemic infections and significant mortality. Pseudomonas
aeruginosa and Staphylococcus aureus are major causal pathogens of endotracheal tube
associated infections with 41.7% and 36.7% VAP cases being attributed to them
respectively [3]. Other microorganisms contributing to VAP are Streptococcus pneumoniae,
Acinetobacter baumannii, Enterobacteriacea and other Gram-negative aerobic bacteria
[2,3].
Many guidelines for the prevention of VAP have been published [12-16], which
including a ventilator bundle launched in 2006 by US agencies and scientific groups [16].
In the following year, guidelines across European were reviewed and several new
guidelines established [13,14]. Among them, the use of endotracheal tubes with subglottic
secretion drainage (SSD) is recommended in all guidelines while tracheostomy is not
endorsed [13-15]. Coating and impregnation of endotracheal tubes with silver fall in the
gray area as these approaches are recommended by some guidelines while some are against
them [13-15]. In 2010, Lorente and co-workers reviewed emerging strategies that had not
been included in the previous guidelines [17]. These strategies include using endotracheal
tubes with ultrathin cuff membranes and SSD, endotracheal tubes with low cuff pressure,
device with balloons to remove biofilm, and saline instillation [17].
Significant research has been devoted to understand the underlying mechanism of
biofilm formation and strategies utilised by these microorganisms to survive antimicrobial
attack [18-21]. With these insights, approaches have been devised which could be
generalized in two categories: 1) inhibition of bacterial colonization [22-28] and 2)
eradication or detachment of formed biofilms [29-33]. As the functional biology of
biofilms is broad and extensive literature is available, this review we will mainly focus on
the emerging strategies for biofilm treatment in these two categories; with emphasis on the
pre-clinical development phase.
2. Biofilm and the mechanism of formation
P. aeruginosa is the most studied biofilm forming bacterial species [34,35]. In simple
terms, biofilms are sessile communities of bacteria, which attach to biotic or abiotic
surfaces in aqueous environments. The cascade of events of biofilm formation are 1)
deposition of a conditioning layer on a surface, 2) transport of planktonic cells to the
surface via diffusive, convective or active flagella-driven transport, 3) initial surface
contact followed by reversible adsorption/desorption of cells on the surface (initial
colonization), 4) irreversible binding onto the surface, rapid propagation and microcolony
formation, 5) matrix production and biofilm maturation; and 6) detachment (release) of
pioneer cells [36]. This process is outlined in Figure 1.
The colonization of bacteria onto surfaces is universally regarded as the initiation of
biofilm communities. At this stage, planktonic cells move along the surface via motility or
Brownian motion. Once the cells are within close proximity of a surface, reversible
colonization occurs when the net attractive forces outweighed the repulsive forces
generated between the bacterial cell and contact surface [37]. This contact results in the
formation of a monolayer of cells on the surface. At this stage, the bacterial cells are still
susceptible to antibiotic treatment.
Based on the Derjaguin-Landau-Verwey-Overbeek (DLVO) adhesion theory, the main
interactions between cells and the surface are through van der Waals and Coulomb forces
[38]. In aqueous conditions, counter ions generated as a result of surface charges are
attracted to each surface to form electric double layers. As bacterial cells are negatively
charged, repulsive electrostatic interactions between them are intensified in surfaces that
having the same net charge as a result of an overlapping electric double layer [39,40].
Additionally, to ensure the bacteria remain irreversible anchored to the solid surface prior
to or after attachment, exopolysaccharides and eDNA surface conditioning materials are
secreted by pioneer cells to act as binding bridges [37]. Finally, bacterial cells can utilize
their pili and flagella to pierce through the potential energy barriers resulting from
repulsive forces owing to their small radii. A study by O’Toole and Kolter showed that the
pili and flagella are necessary for biofilm formation in P. aeruginosa [41]
The next step of biofilm formation involves multiplication and aggregation of attached
monolayer cells into microcolonies. At an appropriate time, these microcolonies
differentiate into mature biofilms which are enveloped within an extrapolymeric substance
(EPS) matrix [42]. EPS comprises polysaccharides (50 to 90% of the organic mass of
biofilm mass), proteins, DNA and lipids [42-44]. EPS are highly hydrated and tenaciously
bound to the surface. Water channels in EPS allow transport of essential nutrients and
oxygen to cells embedded within the biofilm. In vitro, mature biofilms can grow up to 50
μm in thickness with tower-like and mushroom-shaped structures. Bacterial biofilms at this
stage are extremely well-tolerant to antimicrobial agents, and various mechanisms are
activated which include limiting biocide penetration, reduced growth rate and the presence
of persister cells for enhanced survival rate. The final step in biofilm development involves
the dispersion of biofilm in order to liberate and spread bacterial cells to other locations for
initiation of new biofilms.
3. Approaches to prevent biofilm establishment and/or eradication of preformed
biofilms
3.1 Antimicrobial agents
3.1.1 Antibiotics
The inappropriate use of antibiotics is a key reason contributing to the emergence of
antibiotic resistance. To achieve optimal outcome in mechanically ventilated patients with
biofilm infections the appropriate selection of existing antibiotics, duration of treatment,
combination therapy, inhaled antibiotic formulation and development of novel antibiotics
should be considered. The selection of appropriate antibiotics is often difficult mainly due
to the lack of comparative randomized double-blinded studies demonstrating significant
differences between each antibiotic treatment group [45]. Furthermore, some antibiotics
are prone to developing resistance during therapy, which ultimately causes treatment
failure. For example, as bacterial tolerance towards gentamicin during therapy is common,
one attempt to replace gentamicin with amikacin instead was not successful because once
amikacin treatment was discontinued and gentamicin was re-used, the resistance level
returned [46]. Faced with these challenges, a general empiric antibiotic therapy regimen
has been adopted over the years for suspected VAP cases caused by bacterial biofilm
infections. Generally, upon suspicion of P. aeruginosa biofilm occurrence, combination
therapies of anti-pseudomonal antibiotics are used. Commonly, a β-lactam (cephalosporin)
is used in conjunction with a fluoroquinolone (ciprofloxacin or levofloxacin) or an
aminoglycoside (amikacin or tobramycin) [47]. On the other hand, patients with suspected
Gram-positive bacterial infections are often treated with vancomycin as an initial choice
coupled with anti-methicillin resistant Staphyloccus aureus (MRSA) agents such as
linezolid [47]. Recommended therapy for early infections with no risk factors of multidrug
resistant microorganisms is mono-antibiotic therapy with fluoroquinolones, ertapenem,
ceftriaxone or ampicillin [47,48].
In recent years, inhaled antibiotics have been evaluated as a method to prevent biofilm
formation on endotracheal tubes and as a possible adjunctive therapy for VAP [49-53].
Inhalation therapy is associated with high local antibiotic concentrations within the lung,
superior penetrability and much lower systemic toxicity [54]. In a study by Palmer and
co-workers, the concentration of aerosolized antibiotic in sputum was 200-fold higher than
that achieved through systemic administration [55]. To date, the antibiotics used for
inhalation administration include gentamicin, colistin, tobramycin, polymyxin B, amikacin,
ceftazidim and pentamidine [49,51-53,56-60]. Both aerosolized tobramycin and colistin
have been specifically formulated for administration to mechanically ventilated patients
[49-52,57]. Adair and co-workers compared the efficacy of nebulised gentamicin and
parenterally administered cephalosporin to inhibit biofilm formation on endotracheal tubes
[56]. Nebulised gentamicin demonstrated superior performance in terms of higher local
concentration on the tube and controlling bacterial infection compared to parenteral
administration. In addition, biofilm formation was evident on all endotracheal tubes for
patients receiving parenterally administered antibiotic while only 40% of tubes dosed with
nebulized gentamicin showed biofilm formation [56]. Czosnowski and co-workers
demonstrated that inhaled formulation of antibiotic was successful in treating >70% of
ventilator associated infections including infections by multidrug resistant pathogens [61].
In a recent Phase II clinical trial, 29 mechanically ventilated patients with VAP were
randomized to either receive aerosolized amikacin or placebo every 12 h for 7–14 days [62].
Consistent with other findings, delivery of aerosolized amikacin sustained high
concentrations in the lower respiratory tract with negligible adverse effects [62-64]. In
another double-blind, placebo controlled trial with mechanical ventilated patients infected
with Gram-negative pathogens, adjunctive therapy of inhaled amikacin (400 mg) twice per
day in addition to systemic antibiotics helped to substantially reduce systemic antibiotics
administration by 50% on day 7 [63]. Inhaled colistin has been developed as a response to
multidrug resistance in P. aeruginosa and A. baumannii infections in seriously ill
mechanically ventilated patients [49]. Most studies with inhaled colistin provided
encouraging results, with at least 80% cure in patients [57]. Both ciprofloxacin and
aztreonam are currently under evaluation as potential inhaled therapeutic agents. A Phase
III clinical trial for inhaled ciprofloxacin is underway to investigate its effectiveness
towards mechanically ventilated patients [45].
New antibiotics proposed for the treatment of biofilms include drugs such as
tigecycline, ceftobiprole, and telavancin [31,65-68]. Tigecycline is a derivative of
tetracycline with broad spectrum activities against Gram-negative, Gram-positive and
anaerobic strains including MRSA [68]. In a comparison study between antibiotics on their
actions against MRSA biofilm eradication, rifampin, daptomcyin and tigecycline were able
to detach mature biofilms while linezolid, tobramcyin and levofloxacin were only effective
against young biofilms [31]. Recently, tigecycline, a known bacteriostatic agent was
coupled with gentamicin to treat biofilms of clinical MRSA isolates [68]. Both antibiotics
demonstrated synergistic killing towards MRSA biofilms compared to either antibiotic
alone [68]. Telavancin is a new lipoglycopeptide antibiotic which is similar structurally to
vancomycin yet has been modified to incorporate a lipophilic side chain. With such
similarity, televancin is expected to possess potent bactericidal activity towards MRSA.
Recent evidence revealed that telavancin is more effective compared to vancomycin. The
observed MIC for telavancin against MRSA ranged from 0.06 to 1.0 μg/mL while that of
vancomycin was between 0.5 to 2.0 μg/mL [69,70]. With regard to anti-biofilm activity,
telavancin was also consistently more potent than vancomycin [66,67]. For instance the
minimal biofilm eradication concentration for telavancin was 0.125 to 2 μg/mL, while, for
the same action, the required concentration for vancomycin exceeded 512 μg/mL [66].
Other approaches such as the use of electrical and ultrasound therapies to enhance
antibiotic potency have been reported to treat biofilms. Ultrasound therapy is associated
with aiding antibiotic transport through the biofilm or may behave as a stimulus to induce
antibiotic release from coatings [33]. As demonstrated by Norris and co-workers,
formation of P. aeruginosa biofilm was significantly impeded on ciprofloxacin-coated
hydrogels when they were exposed to ultrasound therapy (43 kHz) [33]. Furthermore, by
applying low levels direct currents (70 μA/cm2) in adjuvant with 1.5 μg/mL tobramycin, it
was found that more than 90% of the subpopulation of persister cells in both planktonic and
those embedded in biofilm could be efficiently killed [32]. As such, one could envisage
designs of endotracheal tubes containing antibiotic reservoirs so that, upon infection with a
biofilm, the release of antibiotic could be triggered with external high-intensity ultrasound
[33]. Another interesting idea is to utilize photodynamic therapy to kill bacterial biofilms.
The principle behind this strategy is based upon the generation of reactive oxygen species
by photosensitizing drugs to cause oxidative damage to biofilms. This was evident in a
study in which photosensitizer methylene blue eradicated biofilm in endotracheal tubes
by >99.9% (p < 0.05) after a single treatment [71].
3.1.2. Silver-based compounds
For centuries elemental silver and silver salts have been known for their antimicrobial
activities [72,73]. However, the advent of antibiotics has dramatically reduced the medical
applications of silver. The history of silver as an antimicrobial agent in clinical settings
dates back to 1844 where a German obstetrician used 1% silver nitrate solution to treat
blindness in newborns caused by Postpartum infections [72]. The renewed interest in silver
may be a consequence of the emergence of multiple antibiotic-resistances in bacteria. In
principle, silver-based therapy is advantageous because a) it is effective against various
multidrug resistant microorganisms; b) it causes simultaneous and multiple antibacterial
actions on cells which reduces the chance of cells acquiring resistance and c) it has low
systemic toxicity [72]. Different forms of silver based compounds are used as antimicrobial
therapy, including silver nanoparticles (AgNPs), silver salts, dendrimer-silver complexes
and polymer-silver nanocomposites [29,72-79]. Although the mechanism of silver toxicity
is mainly believed to derive from the release of Ag+ ions, some reports have pointed out
that additional toxicity routes exist with AgNPs.
Recent evidence has demonstrated that in addition to having potent antimicrobial
effects, silver also exerts anti-biofilm activities [29,30,77,78,80-84]. Bjarnshlolt and
co-workers demonstrated that the addition of 5–10 μg/mL silver sulfadiazine was effective
at eradicating established mature P. aeruginosa biofilms while 1 μg/mL had no apparent
effect [78]. In comparison, tobramycin had negligible effect even at high concentrations
(100 μg ml–1). Two interesting findings were noted; a) the tolerance of P. aeruginosa was
dependent on both drug dose and mode of cell growth. Planktonic P. aeruginosa were more
susceptible towards silver sulfadiazine compared to biofilms where a 100-fold higher dose
was required; b) the resistance mechanism of P. aeruginosa biofilms towards silver
sulfadiazine was quorum sensing independent. In other words, the regulation of cell-cell
signaling towards the development of increased antimicrobial resistance was not an
effective protective mechanism against silver since this compound demonstrated no
significant difference in eradicating both quorum sensing activated- and deficient- biofilm
systems [78,84]. Chaudhari and co-workers demonstrated that quorum sensing signal of S.
aureus biofilm communities was quenched in the presence of AgNPs alone but not in the
presence of antibiotic [84]. The surface conditioning layers (e.g., polysaccharides)
deposited by bacteria were thought to be neutralized by AgNPs which thus effectively
hindered biofilm formation [84]. In another study, 100 nM biologically synthesized AgNPs
used against P. aerugionsa and S. epidermis biofilms resulted in ~95% eradication of the
biofilms [75]. The authors argued that the diffusion of AgNPs was not hindered by EPS
matrix as these nanoparticles reach bacterial cells within the biofilm matrix through the
existing water channels, thus were able to impart their antimicrobial activity [75]. Another
type of bio-based AgNPs synthesized by leaf extract broth of Azadirhacta indica showed
uniform spherical particles with average sizes of 50 to 60 nm [85]. These nanoparticles
caused distinct biofilm formation retardation in S. aureus clinical isolates by interrupting
the secretion of carbohydrates and proteins necessary to form the biofilm matrix [85].
The efficacy of AgNPs as anti-biofilm agents varies according to the nanoparticle size,
coating and shape, particle diffusion into the biofilm and type of microorganism
[29,30,77,80,82]. Spherical citrate-capped AgNPs (average mean size of 8 nm) synthesized
via chemical reduction method with sodium borohydride as a reducing agent was used to
establish the correlation between inactivation of planktonic cells and biofilm formation of
P. aeruginosa [80]. Treatment using 10 μg ml–1 of AgNPs hindered more than 60% biofilm
formation even though half of the planktonic cells population survived the treatment.
Complete killing of planktonic cells and inhibition of biofilm formation was not achievable
even at AgNPs concentrations higher than 90 μg ml–1. Although EPS was indeed produced
by these AgNPs-treated cells, the cells had seemingly lost the ability to adhere firmly to
solid surfaces. In addition, it is hypothesized that the presence of AgNPs caused a change
in energy balance, which subsequently activated a stress response in the bacterial cells
[74,80]. Crucial intracellular components such as DNA or ribosomes were found
crystallized in the centre of bacterial cells upon exposure to AgNPs or silver ions [80,86].
Kora and Arunachalam reported that 45-nm sodium dodecyl sulfate (SDS)-capped AgNPs
synthesized by means of UV photoreduction demonstrated superior activity against biofilm
formation. Complete biofilm inhibition was observed at concentration of AgNPs as low as
1 μg ml–1 [77]. It was probable that these SDS-capped AgNPs exerted their toxicity
towards P. aeruginosa through the generation of reactive oxygen species (ROS) radicals
which in turn damaged cell membranes, increased cell permeability and caused leakage of
intracellular contents [77].
It is interesting to consider the net surface charge differences of stabilizers (i.e. SDS,
citrate or polyvinylpyrrlidone PVP) on the efficacy of silver particles (i.e. via retention of
the silver ions or inhibition of local oxidation). The cell wall of Gram-negative bacteria are
comprised of a lipopolysaccharide membrane and inner peptidoglycan layer in which the
amino, carboxyl and phosphate groups on its cellular membrane provide the bacterium
with a negative charge. Citrate is anionic in nature and PVP is cationic while SDS is a
known amphiphilic molecule. Being amphiphilic, SDS-capped AgNPs can be incorporated
easily into the phospholipid bilayer of the bacterial membrane, which increases the
interactions between AgNPs and membrane proteins. In comparison however, there exists
a certain amount of electrostatic repulsion between negatively charged citrate-capped
AgNPs and P. aeruginosa cells, which limits the cell-AgNPs interactions. Furthermore,
Kittler and co-workers showed that the dissolution rate of AgNPs stabilized using a
negatively charged molecule (i.e. citrate) was lower than a positively charged molecule (i.e.
PVP), probably owing to the role of citrate as a chemical barrier, preventing the release of
Ag+ [76].
AgNPs are reported to be lethal to all microorganisms associated within biofilms [87],
though a recent study by Martinez-Gutierrez and co-workers suggested that the
anti-biofilm behavior of these nanoparticles was strain-dependent [30]. Gram-negative
bacteria biofilms (P. aeruginosa, Acinetobacter baumanii) were more susceptible towards
AgNPs compared to Gram-positive staphylococci biofilms [30]. An approximate1-log
reduction was noted for Gram-negative strains compared to <0.5-log reduction for
Gram-positive when 500 mg ml–1 of AgNPs were administered to mature biofilms in vitro
[30]. Interestingly, irrespective of growth media, the biofilm eradication efficiency was
size-dependent; the smaller the AgNP, the higher the amount of biofilm removal [29]. The
treatment of P. aeruginosa biofilm with 600 µg ml–1 followed a decreasing trend: 8-nm
AgNPs (90%) > 20-nm AgNPs (69%) > 35-nm AgNPs (52%) [29]. The presence of
charged organic matter also altered biofilm resistance to both AgNPs and dissolved silver
ions [88]. Stable non-agglomerated AgNPs was found to exert greater toxicity to P.
fluorescens biofilms compared to both highly agglomerated particles and dissolved ions.
Dissolved silver ions were non-toxic to bacterial cells due to the complexation of cationic
Ag+ with anionic biomacromolecules present on the surface of biofilm matrix, thus
reducing the number of available ionized silver for bactericidal action [88]. The authors
suggested that the transport of AgNPs aggregates was hindered by EPS, limiting AgNPs
delivery directly to adherent cells. In comparison, non-aggregated AgNPs could diffuse
into the biofilm matrix easily and exert their toxicity either via direct nanoparticle-cell
interactions or through slow AgNP dissolution. In this way, stable AgNPs remained toxic
to bacterial biofilms and inactivation of ionic compounds (Ag+) with surface EPS is
minimised [88].
3.2 Surface modification and incorporation of antimicrobial agents
In general, the three main factors that influence bacterial adhesion onto solid surfaces
are i) microenvironment (pH, ionic strength, nutrient availability); ii) type and
characteristics of the bacteria; and iii) physicochemical and morphological properties of
the surface (hydrophobicity, surface roughness, energy) [22,28,89-93]. Surface
modification of biomaterials (i.e. endotracheal tubes) to hinder bacterial colonization is an
effective biofilm control strategy (Table 1).
As discussed, mature biofilms are extremely recalcitrant. Therefore, many believe that
delaying or inhibiting bacteria attachment in the first place is a logical strategy.
[22,24-26,28]. To date, strategies for surface modification of medical materials include i)
physical modification to alter the surface topography of materials without addition of
antimicrobial agents; ii) physical deposition using plasma, ion beam or corona discharge;
iii) covalent binding of antimicrobial agents onto surfaces; iv) coating with a low energy
polymer; and v) surface oxidation.
Lopez-Lopez and co-workers evaluated the kinetics of adherence of S. aureus, S.
epidermis and P. aeruginosa on medical plastics made from different materials such as
PVC, Teflon®, siliconised latex, polyurethane and Vialon® [93]. The authors found that
material type was an important parameter for the rate of bacterial colonization whereby
both polyurethane and Vialon® were the most hostile surface for staphylococci attachment
while E. coli and P. aeruginosa tended to adhere the least to Teflon® [93]. Since then,
many studies have revealed various factors are involved in the irreversible attachment of
cells to a surface. Of particular significance in cell attachment is the surface properties
including surface hydrophobicity, surface energy, porosity and chemical composition
[90-92,94].
3.2.1
Hydrophilic vs. hydrophobic
A hydrophobic surface, as possessed by medical polymers, is often thought to be
favored for bacterial colonization. However, conclusive remarks on this topic are difficult,
as many conflicting data on the role of hydrophobicity on bacterial adhesion have been
reported. This is likely to be predominately due to the classification of hydrophobicity,
surface chemistry and nano-macroscopic structure. Many reports have demonstrated
reduced bacterial adhesion to hydrophobic surfaces, however this is often linked to an
increase in contact angle due to nanoscopic roughness (i.e. the lotus leaf affect).
Conversely other studies have shown reduced adhesion to hydrophilic surfaces likely to be
attributed to surface chemistry. This aspect is an important consideration when reviewing
the literature.
To increase the surface hydrophilicity of PVC endotracheal tubes, Balazs and
co-workers employed a chemical surface modification technique via oxygen glow
discharge to introduce oxygenated functional groups on the tube [94]. Oxygen-plasma
treated PVC surfaces became significantly more hydrophilic, as demonstrated by a
decrease in water contact angle from 80° to 8–20°. Meanwhile, these treated surfaces
showed large defects in the order of 15–30 μm, which could be a direct consequence of
incorporating oxygenated functional groups. The authors found that the treated
hydrophobic PVC tubes showed a 70% reduction in adhesion of four different strains of P.
aeruginosa [94]. In another study, the adhesion of eighteen clinical isolates of P.
aeruginosa on oxygen-plasma treated PVC endotracheal tubes was reported [92]. As
expected, a more hydrophilic surface was better at hindering P. aeruginosa adhesion by as
much as 70%. It is noteworthy to mention that the bacteria adhesion behavior differed
significantly between strains. For example, four clinical strains isolated from endotracheal
aspiration of ICU patients adhered at least 600% better to a surface than the model P.
aeruginosa PAO1 laboratory strain, which indicates that future interventional strategies
aimed to reduce bacterial adhesion should be studied on various P. aeruginosa strains
rather than relying on a single laboratory strain [92]. Sousa and co-workers showed that the
affinity of S. epidermis towards acrylic-based material (less hydrophobic) is lower than
silicone (more hydrophobic) [90].
In contrast, Tang and co-workers reported that in vitro bacterial adhesion was reduced
42–89% with a more hydrophobic surface. [91]. However, it is likely that this may be
reflective of the surface roughness rather than chemistry. The susceptibilities of smooth
and nano-rough PVC endotracheal tubes to bacterial adhesion were evaluated in a recent
study [28]. For this, commercially available endotracheal tubes (Sheridan® 6.0 mm ID,
uncuffed) were subjected to lipase digestion to create a nano-rough texture. Using various
validation techniques to determine the textural preferences of P. aeruginosa, it was
concluded that nano-rough PVC endotracheal tubes reduced colonization by about 40%
compared to smooth surfaces [28].
3.2.2. Modification of endotracheal tube surface topography at a nano-scale
Surface roughness at the nano-scale is suggested to minimize contact between the
bacterial cell wall and the surface, thereby reducing the electrostatic interactions for initial
cell attachment [28]. Furthermore, coating of a sugar metabolite (i.e. fructose) onto
nano-rough PVC endotracheal tube surfaces was shown to reduce both planktonic growth
and biofilm formation on treated nano-rough surfaces [26]. Sugar metabolites such as
fructose have also been used in tandem with aminoglycosides to treat persister cells of both
Gram-positive and Gram-negative microorganisms. Fructose is believed to act as
stimulator to switch dormant persister cells into actively growing cells that are more
susceptible to antibiotics [95]. Interestingly, serum was found to actively inhibit the
formation of P. aeruginosa biofilm on medical devices, possibly owing to hindrance of
interaction between microorganisms and the surface or by inhibiting directed bacterial
twitching motility [96].
Inspired by the non-wetting and self-cleaning behavior of the lotus leaf, a recent study
attempted to replicate the characteristics of lotus leafs to achieve an anti-bacterial
anti-adhesion surface [22]. In particular, the treatment of PVC plastics using solvent
(tetrahydrofuran, THF) and anti-solvent (ethanol or methanol) induced morphological
changes in surfaces of endotracheal materials with notable increase in surface
hydrophobicities [22]. The authors showed that ethanol-treated PVC surfaces were
super-hydrophobic containing submicron-textured structures; resulting in significant delay
of P. aeruginosa attachment without using antibiotics. Untreated smooth control PVC
surface was colonized with P. aeruginosa as early as 6 h and initiation of biofilm
maturation was evident at 24 h. On the other hand, bacterial cells were only visible on
treated super-hydrophobic PVC surface at 24 h [22].
Artificial adaptation of shark’s skin has led to design novel biomimetic surfaces
composed of pillars or spikes with varying heights, diameter and space separation [25].
These imprinted micro-patterns of sub-micron pillars are believed to control
microorganism attachment [24,25,97,98] without altering mechanical properties and
compatibility of the bulk material. Chung and co-workers proved that these patterned
surfaces were resistant to S. aureus biofilm formation up to 21 days. Meanwhile, mature
biofilm was initiated on smooth surface at day 7 [97]. A similar approach was employed to
investigate the effects of micro-patterning texture of medical plastics on staphylococci
adhesion and biofilm formation [24]. Using a soft-lithography technique, polyurethane
biomaterial surface was imprinted with ordered and uniformed pillars having an available
(accessible) surface contact area of ~25% compared to smooth and non-textured surface
[24]. Initial adhesion of S. aureus was reduced to 90% thus subsequent biofilm
development was markedly impaired [24]. As these pillars had an average height of 700
nm and the largest pillar space separation was of sub-bacterial dimension, these effectively
restricted the contact of S. aureus to the material surface, leaving the top of the pillars the
only point cells could interact with the surface [24]. A novel biomimetic
submicron-patterned surface in an endotracheal tube-like polymer was engineered recently
to evaluate the effect of patterned topography on S. aureus biofilm formation [25]. The use
of these micro-patterned textures resulted in 89% inhibition when biofilms were grown for
4 days in standard culture containing mucin [25].
3.3.3
Coating an impregnation of endotracheal tubes
Coating of antimicrobial agents using chemical modification techniques has also been
a subject of intense interest lately. Silver-impregnated coating on endotracheal tubes is
probably the most studied with extensive investigations at clinical levels [99-104]. Other
antimicrobial agents currently under investigation in vitro (or pre-clinical) settings include
other silver-based compounds, zinc oxide nanoparticles, thiocyanates, bronopol,
benzalkonium chloride, chlorohexidine, triclosan and hexetidine [23,27,105,106]. The
immobilization of zinc oxide (ZnO) nanoparticles onto PVC endotracheal tubes was found
to retain the bacteriostatic nature of ZnO as the ratio of live to dead S. aureus cells
compared PVC alone [27]. The antibacterial activity was attributed to the release of
bacteriostatic Zn2+ from nanoparticles, bacterial cell membranes binding to ZnO
nanoparticles and formation of reactive oxygen species in cells [27]. On the other hand,
thiocyanation of PVC surfaces, via covalent immobilization, imparted hydrophilicity and
bactericidal characteristics to the treated surface. Although the bactericidal effect of
immobilized thiocyanate was lower than that of free soluble thiocyanate, this did not affect
the anti-adhesion effect of the treated surfaces, which was shown by lower adhesion of
staphylococci in the thiocyanate treated PVC compared to non-treated PVC surfaces [105].
Physical deposition of triclosan and bronopol onto medical grade PVC using a plasma
immersion ion implantation technique was applied to induce antibacterial properties on
PVC surfaces. This approach is primarily based on the coatings of triclosan and bronopol
onto surfaces pre-treated with oxygen plasma, followed by modification of coated
molecules with argon plasma to improve their antibacterial characteristics [23]. From these
results, both triclosan and bronopol were effective against S. aureus with ~80% cells killed
after 10 days of culture but were unable to hinder the colonization of S. aureus [23]. On the
other hand, triclosan-treated surfaces were more effective against E. coli compared to
bronopol. The adherence of survived E. coli cells onto PVC surfaces was also significantly
lower than S. aureus [23]. The deposition of antimicrobial agents onto medical grade PVC
using a multistep physicochemical approach was achieved via surface discharge plasma
and graft copolymerization to produce high-density structures and functionalization with
antimicrobial agents such as bronopol, bezalkonium chloride and chlorohexidine [106].
Both bronopol and bezalkonium chloride were only effective against E. coli (up to 80%
inhibition in adhesion) while the adhesion of S. aureus was not different compared to the
control group. In comparison, the incorporation of chlorohexidine effectively blocked the
attachment of both Gram-positive and Gram-negative bacteria [106].
To-date, coatings of endotracheal tubes with silver-based compounds have received
the most significant attention. Early in vitro, in vivo and pre-clinical trials on the effects of
silver-coated endotracheal tubes on bacterial colonization, and bacterial resistance have
demonstrated promising results [99-103,107]. In 1999, Hartmann and co-workers reported
the first investigation on silver coated endotracheal tubes. The authors used an in vitro
oropharynx-larynx-lung model which was continuously exposed to P. aeruginosa and
mechanically ventilated up to 50 h to closely mimic the actual clinical scenario [107]. This
study found that non-coated control tubes were colonized to a greater extent compared to
silver-coated tubes [107]. A randomized double-blinded controlled experiment using a
mechanically ventilated dog model challenged with buccal administration of P. aeruginosa,
demonstrated that silver-coated endotracheal tubes exerted sustained antimicrobial effect
within the lung airways and hindered biofilm formation on the tubes [102]. Delayed onset
of lung colonization of P. aeruginosa was observed for subjects receiving silver-coated
tubes (1.8 ± 0.8 vs. 3.2 ± 0.8 days, p = 0.02) and the total count of bacterial burden in lung
parenchyma was also significantly reduced (44.8 ± 0.8 vs. 5.4 ± 9 log CFU g–1, p = 0.01)
compared to uncoated tubes [102]. Similarly, Berra and co-workers also performed a
controlled, randomized study in 16 sheep, mechanically ventilated with either control
uncoated endotracheal tubes or silver-sulfadiazine and chlorhexidine coated tubes to
compare the rate of bacterial colonization on the tubes and ventilator circuits after 24 h
[101]. All control tubes were severely colonized (up 108 CFU g–1) and demonstrated thick,
densely packed and aggregated bacterial biofilms (193.3–405.6 μm). The trachea and lungs
in five of eight control groups were infected with pathogenic bacteria. However, bacterial
colonization within coated tubes and the entire ventilator circuits was hindered in seven of
eight ventilated sheep. Biofilm formation on endotracheal tubes was also not evident and
the lungs showed no traces of bacterial colonization [101]. Berra and co-workers evaluated
the feasibility of silver sulfadiazine coated endotracheal tubes challenged with 104–106
CFU ml–1 P. aeruginosa every 24 h in in vitro as well as in animals [100]. In the in vitro
setting, the coated tubes remained bacteria free up to 72 hours while non-coated
endotracheal tubes were heavily colonized with bacteria (up to 3.2 x 109 CFU g–1) [100].
This was further supported in animal studies where thick mucoid biofilm layers were
formed on the non-coated tube, ventilator tubing and lower respiratory tract (p ˂ 0.01). The
common aerobic microorganisms found were α-hemolytic Streptococcus spp., K.
pneumoniae, Moraxella spp., Pasteurella haemolytica, P. multocida, Pseudomonas
aeruginosa and Staphylococcus aureus [100]. The mechanically ventilated sheep with
silver sulfadiazine coated tubes meanwhile were associated with decreased bacterial
burden on the tubes, ventilator circuit and respiratory tract. No local or systemic toxicity
were observed [100]. Similar findings were also recorded in a series of investigations using
in vitro and animal (rabbits) models, [103].
A prospective, randomized phase I–II clinical trial was carried out with forty-six
patients undergoing cardiac surgery to primarily establish the interventions of silver
sulfadiazine coating onto endotracheal tubes to reduce bacterial burden in patients
receiving mechanical ventilation for 12–24 h [99]. The patients, above 18 of age, who
require mechanical ventilation in anesthesia condition were randomized into two groups to
receive either a silver sulfadiazine coated or a conventional standard endotracheal tube [99].
In another independent small clinical trial, the safety of silver-coated endotracheal tubes
and associated bactericidal activity in the lung airways were reported [104]. For this 121
patients who needed mechanical ventilation for more than 24 h and did not have prior
respiratory infections were recruited to randomly receive either silver-coated or non-coated
endotracheal tubes [104]. In this study, Ag+ was dispersed in a polymer on both the inner
and outer lumens and could migrate to the surface of endotracheal tube to provide sustained
bactericidal effect [104]. In general, both studies demonstrated that the use of silver coated
endotracheal tubes was associated with reduced bacterial colonization on the tube lumen
and non-appearance of biofilm formation. Furthermore, these tubes were safely
implemented, easy to manage and well-tolerated with no significant adverse reactions
[99,104]. Furthermore, probably due to the small sample groups, the impact on VAP
incidence was not sufficiently demonstrated [104].
In a large randomized North American silver-coated endotracheal tube (NASCENT)
single-blind trial study, a total of 2003 patients expected to use ventilators for more than 24
h were recruited and randomized to receive either silver-coated or conventional
endotracheal tubes [108]. Their findings could be summarized as follows: a) silver-coated
endotracheal tubes reduced VAP incidence from 7.5% to 4.8% which corresponded to a
relative risk reduction of 35.9% and an absolute risk reduction of 2.7%; b) silver-coated
endotracheal tubes were not successful to achieve significant reduction in mortality rate,
duration of intubation, duration of ICU stay or the severity of adverse effects compared to
conventional tubes [108]. The same group further evaluated mortality in patients who
developed VAP in the previous NASCENT trial [108] using retrospective cohort analysis
[109]. The silver-coated tube was associated with reduced mortality with VAP (14%)
compared to control (36%). However, no differences were observed for those who did not
develop VAP [109]. In addition, the use of silver-coated endotracheal tubes as a
preventative measure for VAP might result in potential hospital cost savings [110].
4. Conclusions
Numerous studies have aimed to investigate the role of endotracheal tubes as
causative agent on VAP as it remains the most commonly acquired infections in intubated
patients. It is generally accepted that rapid formation of multiple-drug-resistant bacterial
biofilms is a main contributor to VAP. The insertion of endotracheal tubes into patients
bypasses the body’s primary host defenses, in which the lung becomes a suitable site for
bacterial colonization, proliferation and biofilm formations. The prevention of VAP or
rather the inhibition of bacterial biofilms on endotracheal tubes is becoming a priority for
hospitals. To date, many strategies have been attempted to eradicate or prevent biofilm
contamination on endotracheal tubes, which include use or biocide, antibiotics and
dispersal agents, ultrasound, chelation, enzymatic digestion or surface modification of
surface. Although these methods are promising, many afford only temporary relief as
microorganisms are quick to develop inherent resistance. The main question is, how much
understanding do we have on the intricate systems of biofilm communities? As these bugs
are constantly evolving, it is desirable to design therapeutic approaches that combine
several modes of antimicrobial actions synergistically to achieve greater sensitivity of
biofilms
5. Expert commentary
Ventilator associated pneumonia in ICU is a significant healthcare problem, affecting up to
28% of intubated patients. The intubation tube provides the first point of call for bacterial
infection, since this invasive medical device is open to bacterial adhesion and biofilm
formation. There is no standardized guidelines for preventing VAP in hospital and a
number of recommendations exist including endotracheal tubes with low cuff pressure,
devices with balloons to remove biofilms, and saline instillation and use of silver. At a
research and development level, a number of stratergies are under development to tackle
bacterial adhesion, biofilm formation and to target bacteria locally, with a view to reduce
and treat VAP. One approach is to modify the surface chemistry or surface topology of the
endotracheal tube to be more hydrophilic or contain nano-rough surfaces. While these may
reduce bacterial adhesion, this is likely to only temporarily prevent adhesion, since bacteria
express and lay down conditioning media that will eventually overcome this approach. A
second approach is to incorporate antibiotics or antimicrobial compounds into/onto the
tube. Approaches such as the incorporating silver nanoparticles may enhance bacterial
killing however; again this approach is likely to be overcome with time. Ultimately, a
multifaceted approach is likely to be the answer to preventing VAP; utilizing tubes that
incorporate both modified surfaces and antibacterials.
6. Five-year view
Currently, the main focus of clinical trials has been to reduce and treat VAP via
impregnation or coating of endotracheal tubes with silver. Research within the field has
mainly focused on two areas, incorporation of antibacterials into/onto tubes or surface
modification to hinder bacteria adhesion and thus biofilm formation. Over the next five
years it is likely that we will see both these novel approaches reach a clinical setting. The
most-likely successful approach, however is to take a multi-faceted approach, via
modification of surface texture and incorporation of antibacterials into the surface structure.
Importantly, however, the success of modified endotracheal tubes in reducing VAP, is
going to come down to cost effectiveness. A successful clinical product is likely to be
cheap to manufacture and not require extensive processing over normal manufacturing
processes. Furthermore, incorporation of currently used antibacterials and modification of
polymers already used in the clinic would ensure rapid translation and uptake in the clinic
if proved effective in preventing VAP.
7. Key issues

Endotracheal tubes in ICU are essential for ventilation but are a major source of
infection and ventilator associated pneumonia (VAP).

VAP affects up to 28% of intubated patients and the incidence rate escalates with
time post ventilation.

After bacterial adhesion to intubation tubes, biofilm formation can occur making it
difficult to remove potential sources of VAP via conventional antibiotics.

Guidelines for reducing VAP are not uniform and there is no cohesive clinical
strategy for reducing VAP associated mortality.

Reducing bacterial adhesion is a key strategy for reducing biofilm formation and
VAP through surface modification of intubation tubes.

Actively ‘targeting bacteria’ by incorporating antibacterial drugs or metal
ions/nanoparticles into tubes is another strategy.

Clinical trials have shown some success using silver coated endotracheal tubes.

Developing improved endotracheal tubes that prevent VAP and reduce VA related
mortality has the potential to improve healthcare outcomes and reduce ICU costs.
Figure captions
Figure 1: Schematic representative of the major stages in the development of P.
aeruginosa biofilm
Figure 2: Proposed biofilm resistance mechanisms. (A) The presence of EPS provides
physical barrier, which limits penetration and diffusion of antimicrobial agents. (B)
Metabolic activities and growth rate of bacterial cells within biofilm matrix as a function of
cell depth is determined by various factors such as nutrient and oxygen availability. The
zone showing the lightest color represents cells with the highest growth rate and hence the
most susceptibility to antimicrobial agents. Cells at the most-inner layers (zone with the
darkest orange color) often survive in dormant state or live in anaerobic conditions. (C)
Activation of stress regulator (RpoS) which could mediate the overexpression of
antimicrobial agent-destroying enzymes. The activation of biofilm resistance genes such as
efflux pumps effectively removes antimicrobial agents from ctyoplasms. (D) Survival of
persister cells which lead to revolution into recurrent ‘superbug’ biofilm infections with
extremely high resistance to antimicrobial agents.
References
* of interest; **of considerable interest.
1.
Chastre J, Fagon J-Y. Ventilator-associated Pneumonia. Am J Respir Crit Care Med,
165, 867-903 (2002).
2.
Koulenti D, Rello J. Hospital-acquired pneumonia in the 21st century: a review of
existing treatment options and their impact on patient care. Expert Opin Pharmaco,
7(12), 1555-1569 (2006).
3.
Ibrahim EH, Tracy L, Hill C, Fraser VJ, Kollef MH. The occurrence of
ventilator-associated pneumonia in a community hospital - risk factors and clinical
outcomes. Chest, 120, 555-561 (2001).
4.
Cook DJ, Walter SD, Cook RJ et al. Incidence of and Risk Factors for
Ventilator-Associated Pneumonia in Critically Ill Patients. Annals of Internal
Medicine, 129(6), 433-440 (1998).
5.
Pneumatikos IA, Dragoumanis CK, Bouros DE. Ventilator-associated Pneumonia
or Endotracheal Tube-associated Pneumonia?: An Approach to the Pathogenesis
and Preventive Strategies Emphasizing the Importance of Endotracheal Tube.
Anesthesiology, 110(3), 673-680 610.1097/ALN.1090b1013e31819868e31819860
(2009).
6.
Sathishkumar S, Fassl J. Endotracheal Tube-associated Pneumonia. Anesthesiology,
111(4), 922 910.1097/ALN.1090b1013e3181b1064c1024 (2009).
7.
Kollef MH. Prevention of hospital-associated pneumonia and ventilator-associated
pneumonia. Crit Care Med, 32(6), 1396-1405 (2004).
8.
Rello J, Soñora R, Jubert P, Artigas A, Rué M, Vallés J. Pneumonia in intubated
patients: role of respiratory airway care. American Journal of Respiratory and
Critical Care Medicine, 154(1), 111-115 (1996).
9.
Levine SA, Niederman MS. The impact of tracheal intubation on host defenses and
risks for nosocomial pneumonia. Clin Chest Med, 12(3), 523-543 (1991).
10.
Feldman C, Kassel M, Cantrell J et al. The presence and sequence of endotracheal
tube colonization in patients undergoing mechanical ventilation. European
Respiratory Journal, 13(3), 546-551 (1999).
11.
Rubenstein JS, Kabat K, Shulman ST, Yogev R. Bacterial and fungal colonization
of endotracheal tubes in children: A prospective study. Crit Care Med, 20(11),
1544-1549 (1992).
12.
Torres A, Carlet J, Bouza E et al. Ventilator-associated pneumonia. European
Respiratory Journal, 17(5), 1034-1045 (2001).
13.
Torres A, Ewig S, Lode H, Carlet J, Grp EHW. Defining, treating and preventing
hospital acquired pneumonia: European perspective. Intens Care Med, 35(1), 9-29
(2009).
14.
Rello J, Lode H, Cornaglia G, Masterton R, Contributors VCB. A European care
bundle for prevention of ventilator-associated pneumonia. Intens Care Med, 36(5),
773-780 (2010).
15.
Muscedere J, Dodek P, Keenan S et al. Comprehensive evidence-based clinical
practice guidelines for ventilator-associated pneumonia: Prevention. J Crit Care,
23(1), 126-137 (2008).
16.
Berwick DM, Calkins DR, McCannon C, Hackbarth AD. The 100 000 lives
campaign: Setting a goal and a deadline for improving health care quality. JAMA,
295(3), 324-327 (2006).
17.
Lorente L, Blot S, Rello J. New Issues and Controversies in the Prevention of
Ventilator-associated Pneumonia. American Journal of Respiratory and Critical
Care Medicine, 182(7), 870-876 (2010).** review of current guidelines
18.
Drenkard E. Antimicrobial resistance of Pseudomonas aeruginosa biofilms.
Microbes Infect, 5(13), 1213-1219 (2003).
19.
Walters MC, Roe F, Bugnicourt A, Franklin MJ, Stewart PS. Contributions of
antibiotic penetration, oxygen limitation, and low metabolic activity to tolerance of
Pseudomonas aeruginosa biofilms to ciprofloxacin and tobramycin. Antimicrob
Agents Ch, 47(1), 317-323 (2003).
20.
Fernández L, Breidenstein EBM, Hancock REW. Creeping baselines and adaptive
resistance to antibiotics. Drug Resistance Updates, 14(1), 1-21 (2011).
21.
Ortíz-Pérez A, Martín-de-Hijas N, Alonso-Rodríguez N, Molina-Manso D,
Fernández-Roblas R, Esteban J. Importance of antibiotic penetration in the
antimicrobial resistance of biofilm formed by non-pigmented rapidly growing
mycobacteria against amikacin, ciprofloxacin and clarithromycin. Enfermedades
Infecciosas Y Microbiologia Clinica, 29(2), 79-84 (2011).
22.
Loo CY, Young PM, Lee WH, Cavaliere R, Whitchurch CB, Rohanizadeh R.
Super-hydrophobic nanotextured polyvinyl chloride films for delaying
Pseudomonas aeruginosa attachment to intubation tubes and medical plastics. Acta
Biomater., 8(5), 1881-1890 (2012).*Example of endotracheal tube material sufrace
modifiaction using a simple process on bacterial adhesion and biofilm formation
23.
Zhang W, Chu PK, Ji J et al. Plasma surface modification of poly vinyl chloride for
improvement of antibacterial properties. Biomaterials, 27(1), 44-51 (2006).
24.
Xu LC, Siedlecki CA. Submicron-textured biomaterial surface reduces
staphylococcal bacterial adhesion and biofilm formation. Acta Biomater, 8(1),
72-81 (2012).
25.
Hoffman M, May RM, Reddy ST. Micro-Patterned Surfaces for Reducing Biofilm
Formation in an Endotracheal-Tube-Like Environment. American journal of
infection control, 40(5), e59-e60 (2012).
26.
Durmus NG, Taylor EN, Inci F, Kummer KM, Tarquinio KM, Webster TJ.
Fructose-enhanced reduction of bacterial growth on nanorough surfaces. Int. J.
Nanomedicine, 7, 537-545 (2012).
27.
Seil JT, Webster TJ. Reduced Staphylococcus aureus proliferation and biofilm
formation on zinc oxide nanoparticle PVC composite surfaces. Acta Biomater, 7(6),
2579-2584 (2011).
28.
Seil JT, Rubien NM, Webster TJ, Tarquinio KM. Comparison of quantification
methods illustrates reduced Pseudomonas aeruginosa activity on nanorough
polyvinyl chloride. Journal of Biomedical Materials Research Part B: Applied
Biomaterials, 98B(1), 1-7 (2011).
29.
Loo CY, Young PM, Cavaliere R, Whitchurch CB, Lee WH, Rohanizadeh R. Silver
nanoparticles enhance Pseudomonas aeruginosa PAO1 biofilm detachment. Drug
Dev. Ind. Pharm., Early online, 1-11 (2013).*Effect of silver nanoparticles and free
silver nitrate on antibacterial effect for VAP
30.
Martinez-Gutierrez F, Boegli L, Agostinho A et al. Anti-biofilm activity of silver
nanoparticles against different microorganisms. Biofouling, 29(6), 651-660 (2013).
31.
Cafiso V, Bertuccio T, Spina D, Purrello S, Stefani S. Tigecycline inhibition of a
mature biofilm in clinical isolates of Staphylococcus aureus : comparison with
other drugs. FEMS Immunology & Medical Microbiology, 59(3), 466-469 (2010).
32.
Niepa THR, Gilbert JL, Ren DC. Controlling Pseudomonas aeruginosa persister
cells by weak electrochemical currents and synergistic effects with tobramycin.
Biomaterials, 33(30), 7356-7365 (2012).
33.
Norris P, Noble M, Francolini I et al. Ultrasonically controlled release of
ciprofloxacin from self-assembled coatings on poly(2-hydroxyethyl methacrylate)
hydrogels for Pseudomonas aeruginosa biofilm prevention. Antimicrob Agents Ch,
49(10), 4272-4279 (2005).
34.
O'Toole GA. To build a biofilm. J Bacteriol, 185(9), 2687-2689 (2003).*
Formation of biofilms; background information
35.
Stover CK, Pham XQ, Erwin AL et al. Complete genome sequence of
Pseudomonas aeruginosa PAO1, an opportunistic pathogen. Nature, 406(6799),
959-964 (2000).
36.
Palmer RJ, White DC. Developmental biology of biofilms: implications for
treatment and control. Trends Microbiol., 5(11), 435-440 (1997). * Formation of
biofilms; background information
37.
Hori K, Matsumoto S. Bacterial adhesion: From mechanism to control. Biochem
Eng J, 48(3), 424-434 (2010).
38.
Hiemenz PC, Rajagopalan R. Principles in colloid and surface chemistry (Decker,
M., New York, 1997).
39.
Loosdrecht MM, Lyklema J, Norde W, Zehnder AB. Bacterial adhesion: A
physicochemical approach. Microb Ecol, 17(1), 1-15 (1989).
40.
Ong YL, Razatos A, Georgiou G, Sharma MM. Adhesion forces between E-coli
bacteria and biomaterial surfaces. Langmuir, 15(8), 2719-2725 (1999).
41.
O'Toole GA, Kolter R. Flagellar and twitching motility are necessary for
Pseudomonas aeruginosa biofilm development. Mol Microbiol, 30(2), 295-304
(1998).
42.
Tsuneda S, Aikawa H, Hayashi H, Yuasa A, Hirata A. Extracellular polymeric
substances responsible for bacterial adhesion onto solid surface. Fems Microbiol
Lett, 223(2), 287-292 (2003).
43.
Flemming HC, Wingender J. Relevance of microbial extracellular polymeric
substances (EPSs) - Part II: Technical aspcets. (Ed.^(Eds) (2001) 9-16.
44.
Flemming HC, Wingender J. Relevance of microbial extracellular polymeric
substances (EPSs) - Part I: Structural and ecological aspects. (Ed.^(Eds) (2001) 1-8.
45.
El Solh AA, Alhajhusain A. Update on the treatment of Pseudomonas aeruginosa
pneumonia. J Antimicrob Chemoth, 64(2), 229-238 (2009).
46.
Gerding DN, Larson TA, Hughes RA, Weiler M, Shanholtzer C, Peterson LR.
Aminoglycoside Resistance and Aminoglycoside Usage - 10 Years of Experience
in One Hospital. Antimicrob Agents Ch, 35(7), 1284-1290 (1991).
47.
Guidelines for the Management of Adults with Hospital-acquired,
Ventilator-associated, and Healthcare-associated Pneumonia. American Journal of
Respiratory and Critical Care Medicine, 171(4), 388-416 (2005).
48.
Niederman MS, Craven DE. Guidelines for the Management of Adults with
Hospital-acquired, Ventilator-associated, and Healthcare-associated Pneumonia.
Am. J. Respir. Crit. Care Med., 171(4), 388-416 (2005).
49.
Korbila IP, Michalopoulos A, Rafailidis PI, Nikita D, Samonis G, Falagas ME.
Inhaled colistin as adjunctive therapy to intravenous colistin for the treatment of
microbiologically documented ventilator-associated pneumonia: a comparative
cohort study. Clinical Microbiology and Infection, 16(8), 1230-1236 (2010).
50.
Kofteridis DP, Alexopoulou C, Valachis A et al. Aerosolized plus Intravenous
Colistin versus Intravenous Colistin Alone for the Treatment of
Ventilator-Associated Pneumonia: A Matched Case-Control Study. Clinical
Infectious Diseases, 51(11), 1238-1244 (2010).
51.
Griese M, Eismann C, Börner G et al. A Pharmacokinetics and Safety Comparison
of a Highly Concentrated Tobramycin Solution with TOBI. Journal of Aerosol
Medicine and Pulmonary Drug Delivery, (2013).
52.
Hallal A, Cohn SM, Namias N et al. Aerosolized Tobramycin in The Treatment of
Ventilator-Associated Pneumonia: A Pilot Study. Surgical Infections, 8(1), 73-82
(2007).
53.
Lu Q, Yang J, Liu Z, Gutierrez C, Aymard G, Rouby J-J. Nebulized Ceftazidime
and Amikacin in Ventilator-associated Pneumonia Caused by Pseudomonas
aeruginosa. American Journal of Respiratory and Critical Care Medicine, 184(1),
106-115 (2011).
54.
Abu-Salah T, Dhand R. Inhaled Antibiotic Therapy for Ventilator-Associated
Tracheobronchitis and Ventilator-Associated Pneumonia: an Update. Adv Ther,
28(9), 728-747 (2011).
55.
Palmer LB, Smaldone GC, Chen JJ et al. Aerosolized antibiotics and
ventilator-associated tracheobronchitis in the intensive care unit. Crit Care Med,
36(7), 2008-2013 (2008).
56.
Adair CG, Gorman SP, Byers LM et al. Eradication of endotracheal tube biofilm by
nebulised gentamicin. Intens Care Med, 28(4), 426-431 (2002).
57.
Kang C-H, Tsai C-M, Wu T-H et al. Colistin inhalation monotherapy for
ventilator-associated pneumonia of Acinetobacter baumannii in prematurity.
Pediatr Pulm, n/a-n/a (2013).
58.
Le Le J, Ashley ED, Neuhauser MM et al. Consensus Summary of Aerosolized
Antimicrobial Agents: Application of Guideline Criteria. Pharmacotherapy: The
Journal of Human Pharmacology and Drug Therapy, 30(6), 562-584 (2010).
59.
Mohr AM, Sifri ZC, Horng HS et al. Use of Aerosolized Aminoglycosides in the
Treatment of Gram-Negative Ventilator-Associated Pneumonia. Surgical Infections,
8(3), 349-358 (2007).
60.
Claridge JA, Edwards NM, Swanson J et al. Aerosolized Ceftazidime Prophylaxis
against Ventilator-Associated Pneumonia in High-Risk Trauma Patients: Results of
A Double-Blind Randomized Study. Surgical Infections, 8(1), 83-90 (2007).
61.
Czosnowski QA, Wood GC, Magnotti LJ et al. Adjunctive Aerosolized Antibiotics
for Treatment of Ventilator-Associated Pneumonia. Pharmacotherapy, 29(9),
1054-1060 (2009).
62.
Luyt C-E, Clavel M, Guntupalli K et al. Pharmacokinetics and lung delivery of
PDDS-aerosolized amikacin (NKTR-061) in intubated and mechanically ventilated
patients with nosocomial pneumonia. Crit Care, 13(6), 1-10 (2009).
63.
Niederman M, Chastre J, Corkery K, Fink J, Luyt C-E, García M. BAY41-6551
achieves bactericidal tracheal aspirate amikacin concentrations in mechanically
ventilated patients with Gram-negative pneumonia. Intens Care Med, 38(2),
263-271 (2012).
64.
Luyt C-E, Eldon MA, Stass H, Gribben D, Corkery K, Chastre J. Pharmacokinetics
and Tolerability of Amikacin Administered as BAY41-6551 Aerosol in
Mechanically Ventilated Patients with Gram-Negative Pneumonia and Acute Renal
Failure. Journal of Aerosol Medicine and Pulmonary Drug Delivery, 24(4),
183-190 (2011).
65.
Gander S, Kinnaird A, Finch R. Telavancin: in vitro activity against staphylococci
in a biofilm model. J Antimicrob Chemoth, 56(2), 337-343 (2005).
66.
LaPlante KL, Mermel LA. In Vitro Activities of Telavancin and Vancomycin
against Biofilm-Producing Staphylococcus aureus, S. epidermidis, and
Enterococcus faecalis Strains. Antimicrob Agents Ch, 53(7), 3166-3169 (2009).
67.
Smith K, Gemmell CG, Lang S. Telavancin shows superior activity to vancomycin
with multidrug-resistant Staphylococcus aureus in a range of in vitro biofilm
models. Eur J Clin Microbiol Infect Dis, 1-6 (2013).
68.
McConeghy KW, LaPlante KL. In vitro activity of tigecycline in combination with
gentamicin against biofilm-forming Staphylococcus aureus. Diagnostic
Microbiology and Infectious Disease, 68(1), 1-6 (2010).
69.
Draghi DC, Benton BM, Krause KM, Thornsberry C, Pillar C, Sahm DF.
Comparative surveillance study of telavancin activity against recently collected
gram-positive clinical isolates from across the United States. Antimicrob Agents Ch,
52(7), 2383-2388 (2008).
70.
Krause KM, Renelli M, Difuntorum S, Wu TX, Debabov DV, Benton BM. In vitro
activity of telavancin against resistant gram-positive bacteria. Antimicrob Agents
Ch, 52(7), 2647-2652 (2008).
71.
Biel MA, Sievert C, Usacheva M et al. Reduction of Endotracheal Tube Biofilms
Using Antimicrobial Photodynamic Therapy. Laser Surg Med, 43(7), 586-590
(2011).
72.
Jain J, Arora S, Rajwade JM, Omray P, Khandelwal S, Paknikar KM. Silver
Nanoparticles in Therapeutics: Development of an Antimicrobial Gel Formulation
for Topical Use. Mol Pharmaceut, 6(5), 1388-1401 (2009).
73.
Dallas P, Sharma VK, Zboril R. Silver polymeric nanocomposites as advanced
antimicrobial agents: Classification, synthetic paths, applications, and perspectives.
Adv Colloid Interfac, 166(1-2), 119-135 (2011).
74.
Roe D, Karandikar B, Bonn-Savage N, Gibbins B, Roullet JB. Antimicrobial
surface functionalization of plastic catheters by silver nanoparticles. J Antimicrob
Chemoth, 61(4), 869-876 (2008).
75.
Kalishwaralal K, BarathManiKanth S, Pandian SRK, Deepak V, Gurunathan S.
Silver nanoparticles impede the biofilm formation by Pseudomonas aeruginosa and
Staphylococcus epidermidis. Colloids Surf B, 79(2), 340-344 (2010).
76.
Kittler S, Greulich C, Diendorf J, Koller M, Epple M. Toxicity of Silver
Nanoparticles Increases during Storage Because of Slow Dissolution under Release
of Silver Ions. Chem Mater, 22(16), 4548-4554 (2010).
77.
Kora AJ, Arunachalam J. Assessment of antibacterial activity of silver
nanoparticles on Pseudomonas aeruginosa and its mechanism of action. World J
Microb Biot, 27(5), 1209-1216 (2011).
78.
Bjarnsholt T, Kirketerp-MØLler K, Kristiansen S et al. Silver against Pseudomonas
aeruginosa biofilms. Apmis, 115(8), 921-928 (2007).
79.
Melaiye A, Youngs WJ. Silver and its application as an antimicrobial agent. Expert
Opin Ther Pat, 15(2), 125-130 (2005).
80.
Dror-Ehre A, Adin A, Markovich G, Mamane H. Control of biofilm formation in
water using molecularly capped silver nanoparticles. Water Res, 44(8), 2601-2609
(2010).
81.
Park H-J, Park S, Roh J et al. Biofilm-inactivating activity of silver nanoparticles:
A comparison with silver ions. Journal of Industrial and Engineering Chemistry,
19(2), 614-619 (2013).
82.
Fabrega J, Renshaw JC, Lead JR. Interactions of Silver Nanoparticles with
Pseudomonas putida Biofilms. Environ Sci Technol, 43(23), 9004-9009 (2009).
83.
Leid JG, Ditto AJ, Knapp A et al. In vitro antimicrobial studies of silver carbene
complexes: activity of free and nanoparticle carbene formulations against clinical
isolates of pathogenic bacteria. J Antimicrob Chemoth, 67(1), 138-148 (2012).
84.
Chaudhari PR, Masurkar SA, Shidore VB, Kamble SP. Effect of Biosynthesized
Silver Nanoparticles on Staphylococcus aureus Biofilm Quenching and Prevention
of Biofilm Formation. Nano-Micro Lett, 4(1), 34-39 (2012).
85.
Namasivayam SKR, Christo BB, Arasu SMK, Kumar KAM, Deepak K. Anti
biofilm effect of biogenic silver nanoparticles coated medical devices against
biofilm of clinical isolate of Staphylococcus aureus. Global Journal of Medical
Research, 13(3) (2013).
86.
Feng QL, Wu J, Chen GQ, Cui FZ, Kim TN, Kim JO. A mechanistic study of the
antibacterial effect of silver ions on Escherichia coli and Staphylococcus aureus. J
Biomed Mater Res, 52(4), 662-668 (2000).
87.
Fayaz AM, Balaji K, Girilal M, Yadav R, Kalaichelvan PT, Venketesan R. Biogenic
synthesis of silver nanoparticles and their synergistic effect with antibiotics: a study
against gram-positive and gram-negative bacteria. Nanomed-Nanotechnol, 6(1),
103-109 (2010).
88.
Wirth SM, Lowry GV, Tilton RD. Natural Organic Matter Alters Biofilm Tolerance
to Silver Nanoparticles and Dissolved Silver. Environ Sci Technol, 46(22),
12687-12696 (2012).
89.
Quirynen M, Van Der Mei HC, Bollen CML et al. An in vivo Study of the Influence
of the Surface Roughness of Implants on the Microbiology of Supra- and
Subgingival Plaque. Journal of Dental Research, 72(9), 1304-1309 (1993).
90.
Sousa C, #225, udia, Teixeira P, Oliveira R, rio. Influence of Surface Properties on
the Adhesion of Staphylococcus epidermidis to Acrylic and Silicone. International
Journal of Biomaterials, 2009 (2009).
91.
Tang H, Cao T, Liang X et al. Influence of silicone surface roughness and
hydrophobicity on adhesion and colonization of Staphylococcus epidermidis.
Journal of Biomedical Materials Research Part A, 88A(2), 454-463 (2009).
92.
Triandafillu K, Balazs DJ, Aronsson BO et al. Adhesion of Pseudomonas
aeruginosa strains to untreated and oxygen-plasma treated poly(vinyl chloride)
(PVC) from endotracheal intubation devices. Biomaterials, 24(8), 1507-1518
(2003).
93.
Lopez-Lopez G, Pascual A, Perea EJ. Effect of plastic catheter material on bacterial
adherence and viability. Journal of Medical Microbiology, 34(6), 349-353 (1991).
94.
Balazs DJ, Triandafillu K, Chevolot Y et al. Surface modification of PVC
endotracheal tubes by oxygen glow discharge to reduce bacterial adhesion. Surface
and Interface Analysis, 35(3), 301-309 (2003).
95.
Allison KR, Brynildsen MP, Collins JJ. Metabolite-enabled eradication of bacterial
persisters by aminoglycosides. Nature, 473(7346), 216-220 (2011).
96.
Hammond A, Dertien J, Colmer-Hamood JA, Griswold JA, Hamood AN. Serum
Inhibits P. aeruginosa Biofilm Formation on Plastic Surfaces and Intravenous
Catheters. J Surg Res, 159(2), 735-746 (2010).
97.
Chung KK, Schumacher JF, Sampson EM, Burne RA, Antonelli PJ, Brennana AB.
Impact of engineered surface microtopography on biofilm formation of
Staphylococcus aureus. Biointerphases, 2(2), 89-94 (2007).
98.
May R, Hoffman M, Reddy S. 12: Micro-Patterned Surfaces for Reducing Bacterial
Biofilm Formation on Endotracheal Tubes: A Novel Approach to Decreasing the
Incidence of Ventilator-Associated Pneumonia. Crit Care Med, 40(12), 1-328
310.1097/1001.ccm.0000424268.0000420384.0000424267b (2012).
99.
Berra L, Kolobow T, Laquerriere P et al. Internally coated endotracheal tubes with
silver sulfadiazine in polyurethane to prevent bacterial colonization: a clinical trial.
Intens Care Med, 34(6), 1030-1037 (2008).
100.
Berra L, Curto F, Li Bassi G et al. Antimicrobial-coated endotracheal tubes: an
experimental study. Intens Care Med, 34(6), 1020-1029 (2008).
101.
Berra L, De Marchi L, Yu Z-X, Laquerriere P, Baccarelli A, Kolobow T.
Endotracheal Tubes Coated with Antiseptics Decrease Bacterial Colonization of
the Ventilator Circuits, Lungs, and Endotracheal Tube. Anesthesiology, 100(6),
1446-1456 (2004).
102.
Olson ME, Harmon BG, Kollef MH. SIlver-coated endotracheal tubes associated
with reduced bacterial burden in the lungs of mechanically ventilated dogs*.
CHEST Journal, 121(3), 863-870 (2002).
103.
Rello J, Afessa B, Anzueto A et al. Activity of a silver-coated endotracheal tube in
preclinical models of ventilator-associated pneumonia and a study after extubation
*. Crit Care Med, 38(4), 1135-1140
1110.1097/CCM.1130b1013e3181cd1112b1138 (2010).
104.
Rello J, Kollef M, Diaz E et al. Reduced burden of bacterial airway colonization
with a novel silver-coated endotracheal tube in a randomized multiple-center
feasibility study *. Crit Care Med, 34(11), 2766-2772
2710.1097/2701.CCM.0000242154.0000249632.B0000242150 (2006).
105.
James NR, Jayakrishnan A. Surface thiocyanation of plasticized poly(vinyl
chloride) and its effect on bacterial adhesion. Biomaterials, 24(13), 2205-2212
(2003).
106.
Asadinezhad A, Novák I, Lehocký M et al. An in vitro bacterial adhesion
assessment of surface-modified medical-grade PVC. Colloids and Surfaces B:
Biointerfaces, 77(2), 246-256 (2010).
107.
Hartmann M, Guttmann J, Müller B, Hallmann T, Geiger K. Reduction of the
bacterial load by the silver-coated endotracheal tube (SCET), a laboratory
investigation. Technology and Health Care, 7(5), 359-370 (1999).
108.
Kollef MH, Afessa B, Anzueto A, et al. Silver-coated endotracheal tubes and
incidence of ventilator-associated pneumonia: The nascent randomized trial. JAMA,
300(7), 805-813 (2008).**Large single blind trial of silver coated modified tubes in
intubated patients
109.
Afessa B, Shorr AF, Anzueto AR, Craven DE, Schinner R, Kollef MH. ASsociation
between a silver-coated endotracheal tube and reduced mortality in patients with
ventilator-associated pneumonia. CHEST Journal, 137(5), 1015-1021 (2010).
110.
Andrew F. Shorr MM, Marya D. Zilberberg MM, Marin Kollef MD.
Cost‐ Effectiveness Analysis of a Silver‐ Coated Endotracheal Tube to Reduce
the Incidence of Ventilator‐ Associated Pneumonia • Infection Control and
Hospital Epidemiology, 30(8), 759-763 (2009).*importance of cost effectiveness in
development of ICU endotracheal tubes