Antimicrobial surface functionalization of plastic catheters
by silver nanoparticles
David Roe1, Balu Karandikar1, Nathan Bonn-Savage1, Bruce Gibbins1and Jean-Baptiste Roullet2*
1AcryMed, Inc., 9560 SW Nimbus Avenue, Beaverton, OR 97008, USA;2Oregon Health & Science University,
707 SW Gaines Road, Portland, OR 97221, USA
Received 13 April 2007; returned 19 August 2007; revised 2 January 2008; accepted 11 January 2008
Objectives: To test the antimicrobial activity and evaluate the risk of systemic toxicity of novel
catheters coated with silver nanoparticles.
Methods: Catheters were coated with silver using AgNO3, a surfactant and N,N,N0,N0-tetramethylethy-
lenediamine as a reducing agent. Particle size was determined by electron microscopy. Silver release
from the catheters was determined in vitro and in vivo using radioactive silver (110mAg1). Activity on
microbial growth and biofilm formation was evaluated against pathogens most commonly involved in
catheter-related infections, and the risk for systemic toxicity was estimated by measuring silver biodis-
tribution in mice implanted subcutaneously with110mAg1-coated catheters.
Results: The coating method yielded a thin (?100 nm) layer of nanoparticles of silver on the surface of
the catheters. Variations in AgNO3concentration translated into proportional changes in silver coating
(from 0.1 to 30 mg/cm2). Sustained release of silver was demonstrated over a period of 10 days. Coated
catheters showed significant in vitro antimicrobial activity and prevented biofilm formation using
Pseudomonas aeruginosa and Candida albicans. Approximately 15% of the coated silver eluted from
the catheters in 10 days in vivo, with predominant excretion in faeces (8%), accumulation at the implan-
tation site (3%) and no organ accumulation (? ? 0.1%).
Conclusions: A method to coat plastic catheters with bioactive silver nanoparticles was developed.
These catheters are non-toxic and are capable of targeted and sustained release of silver at the implan-
tation site. Because of their demonstrated antimicrobial properties, they may be useful in reducing the
risk of infectious complications in patients with indwelling catheters.
Keywords: nanotechnology, nosocomial infections, biofilms, biodistribution, mice
More than 200 000 nosocomial bloodstream infections occur
each year in the USA and most of them are related to the use of
intravascular devices.1,2Central venous catheters are a particu-
larly high risk category of devices.3,4According to recent esti-
mates, the use of 1 in 20 of the 7 million central venous
catheters inserted annually is associated with catheter-related
bloodstream infection.2,5Chronic indwelling urinary catheters
also increase the risk of infection, accounting for ?80% of all
nosocomial urinary tract infections.6
The insertion of catheters under sterile conditions is the most
effective measure to prevent catheter-associated infective com-
plications.7Despite concerns that they would lead to compla-
cency regarding septic techniques, catheters with antimicrobial
properties have nonetheless been proposed as a means to
provide additional protection and further reduce the risk of
infection.8,9A number of such devices have been developed10–
*Corresponding author. Tel: þ1-503-494-4979; Fax: þ1-503-494-2781; E-mail: firstname.lastname@example.org
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version of this article for non-commercial purposes provided that: the original authorship is properly and fully attributed; the Journal and Oxford University
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Journal of Antimicrobial Chemotherapy (2008) 61, 869–876
Advance Access publication 27 February 2008
by guest on June 9, 2013
12and tested with variable success in clinical studies.4,6
However, the ideal catheter, a catheter that combines low-cost
coating technology, wide-spectrum and long-lasting antimicro-
bial properties, and safe utilization, has yet to be developed.
The objective of the studies presented in this report was to
develop a new method to deposit silver nanoparticles at low
temperature on plastic catheters, determine the antimicrobial
activity of these catheters in vitro and examine their silver-
releasing properties both in vitro and in vivo. The data suggest
that plastic catheters coated with silver nanoparticles may be
effective in reducing the infectious risk associated with chronic
catheterization in humans.
Materials and methods
Materials and reagents
All chemicals were purchased from Sigma (St Louis, MO, USA).
(0.53 mm internal diameter, 0.94 mm outer diameter; surface ¼
0.463 cm2/cm) for all experiments. Radioactive silver nitrate
(110mAgNO3, spec. act. 22.2 Bq/mg) was obtained from the Oregon
State University Radiation Center (Corvallis, OR, USA).
Silver coating of the catheters
Catheters were coated on both sides with silver using AcryMed
SilvaGard technology (published US patent application 2007/
0003603). In brief, an aqueous solution containing 5.6 g/L Tween
20, 4 mM sodium saccharine and 1–5 mM silver nitrate was first
preparedandstirred for10 min
N,N,N0,N0-tetramethylethylenediamine (0.33%, v/v). The solution
was then placed in a microwave oven, heated to 568C and then
poured over pre-cut, 2 cm long catheter segments placed in screw-
cap tubes. The tubes were centrifuged to remove the air trapped
inside the catheters and quickly placed in a thermostated oven set at
538C. After 16 h, the coating solution was removed and the cath-
eters were rinsed once with each of the following: aqueous solution
of Tween 20 (1.25 mM), 2% HNO3, H2O and isopropanol. After the
isopropanol wash, the catheters were placed on bench-top paper,
dried and sterilized (autoclave at 1218C for 15 min). The same
method was used to prepare catheters coated with radioactive silver,
simply replacing AgNO3 with
were not sterilized.
Silver concentration was determined in randomly selected cath-
eters by absorption spectrophotometry (‘cold’ silver coating) or by
gamma counting (‘hot’ silver coating). Radioactivity was measured
using a Beckman 5500 gamma counter. All counts were corrected
for background radioactivity, counting efficiency and decay using a
half-life of 256 days.
110mAgNO3. Radioactive catheters
Silver nanoparticle analysis
Silver coating was examined by electron microscopy. For particle
size analysis, a drop of silver nanoparticles prepared as described
above was deposited at the surface of a carbon-coated copper grid
and analysed using a Zeiss DSM-960 equipped with a Link
Analytical multi-window, X-ray detector with a no-window capa-
bility. For coating thickness analysis, thin sections of silver-coated
polycarbonate matrix were prepared and viewed by non-disruptive
electron transmission microscopy using a dual beam STRATA 400S
electron microscope (FEI Company, Hillsboro, OR, USA).
Biofilm formation inhibition assay and bactericidal activity
Silver-coated catheters (650 mg silver/g) were placed in tubes (8 ?
1 cm strips/tube) containing growth medium (1 mL/tube; 1% bovine
serum albumin, 0.1% neopeptone, 0.25% glucose) and 103test
microorganisms (test tubes). Six different microorganisms (clinical
strains) were tested: (i) Escherichia coli; (ii) Enterococcus;
(v) coagulase-negative staphylococci; and (vi) Candida albicans.
Tubes containing non-coated catheters and tubes containing growth
medium but no catheter and no microorganism (controls) were pre-
pared in parallel. The tubes were then incubated for either 24, 48 or
72 h at 358C. A count of free floating viable bacteria and the deter-
mination of viable sessile bacteria by means of a biofilm formation
assay were performed at each time point. Biofilm formation was
estimated using the tetrazolium salt XTT assay as previously
described.13,14Results were expressed in per cent inhibition of
growth of free floating organisms or biofilm formation in each test
tube relative to the average growth or biofilm formation in control
tubes (tubes with non-treated catheters).
In vitro silver release procedure
Catheters were coated with radioactive silver and placed in 5 mL
plastic tubes (33 segments of ?2 cm/tube; n ¼ 5 tubes for each type
of catheters). Each tube was filled with 2 mL of saline (enough to
completely immerse the catheters), capped, assayed for radioactivity
and placed in an oven set at 378C (day 0). Every day for 10 days,
the saline solution was collected, assayed for radioactivity and
replaced with fresh solution after brief washing of the catheters.
Silver release was expressed in mg/day.
In vivo silver release and biodistribution
In vivo studies were performed with C57Bl/6J male mice. On the
day of surgery, the animals were weighed and anaesthetized with an
isoflurane–oxygen mixture (4% induction, 2% maintenance). A sub-
cutaneous pocket (2 cm ? 1 cm) was created on the dorsum of each
animal, ?1–2 cm from the nape of the neck. Catheters (14 seg-
ments, 2 cm in length each; total catheter length ¼ 28 cm) coated
with110mAg were placed in the pocket. The pocket was sealed with
one suture and a drop of tissue glue (3M VetbondTM). The animals
were allowed to recover from anaesthesia and then placed into indi-
vidual metabolic cages. They had free access to food and water and
were housed with a 12 h dark–light cycle at 208C. Urine and faeces
were collected every day in the morning, between 8:00 and 9:00
am. Urine and faeces samples were transferred in counting tubes
and assayed for radioactivity on the same day. On day 10, the
animals were weighed, anaesthetized and sacrificed by cervical dis-
location. Organ and tissue collection was then performed followed
by radioactivity measurement of the collected samples. All
procedures were approved by the institutional (OHSU) Ethics
Silver coating of the catheters
The coating method produced catheters coated with a reproduci-
ble and predictable amount of silver. In these experiments,
batches of 35 catheter segments (each ?2 cm in length) were
Roe et al.
by guest on June 9, 2013
coated using initial110mAgNO3concentrations ranging from 1.0
to 5.0 mM (n ¼ 6 batches/given AgNO3concentration; five con-
centrations tested). As shown in Figure 1, the coating was pro-
portional to the initial concentration of AgNO3and was highly
reproducible from batch to batch (coefficient of variation was
0.7+0.1%, mean+SEM, n ¼ 5). The relationship between
temperature and coating was not fully characterized, but prelimi-
nary experiments suggest a quadratic behaviour of the relation-
ship between 40 and 708C, with higher temperatures resulting in
higher silver concentrations deposited at the surface of the cath-
eters (data not shown).
Silver deposition at the surface of the catheters was analysed
by electron microscopy in parallel experiments performed with
non-radioactive silver. In one experiment, a drop of a solution of
silver nanoparticles prepared as described in the Materials and
methods section was deposited at the surface of a carbon-coated
copper grid and analysed by X-ray electron microscopy.
Elemental composition of treated catheters showed peaks due to
Ag in addition to peaks due to C, N and O, the elements making
up the base polymer grid. As illustrated in Figure 2, distinct and
round particles of silver were observed, with diameters ranging
from ?3 to 18 nm (median ¼ 10.7 nm; n ¼ 100) (Figure 2a
and b). Particle density was estimated at 104/mm2on average.
Particles showed typical polyhedral structures indicating twin
boundaries15,16(Figure 2c). Analysis of silver coating could not
be obtained using silver-coated catheter sections as the base
polymer softened during exposure to the electron beam. The
thickness of the coating was thus examined using non-disruptive
electron microscopy and thin (100–150 nm) sections of a poly-
carbonate matrix coated using conditions similar to those
described for catheters. As illustrated in Figure 2(d), the coating
was only 80–120 nm thick, equivalent to ?4–6 layers of silver
Bactericidal activity and biofilm formation
These experiments were performed using catheters coated with
600 mg of non-radioactive silver. The catheters demonstrated
significant antimicrobial activity against all tested microorgan-
isms. They inhibited both cell growth and biofilm formation for
at least 72 h (Table 1). The effect on growth was complete
growth inhibition of all microorganisms but P. aeruginosa (67%
growth inhibition at t ¼ 72 h). The inhibition of biofilm
formation was almost complete for E. coli, S. aureus and
C. albicans, and reached more than 50% for Enterococcus,
coagulase-negative staphylococci and P. aeruginosa after 72 h.
Silver release from coated catheters (in vitro studies)
These experiments were performed with catheters coated with
two concentrations of silver: 600 mg/g of catheter, a con-
centration similar to that used in the antimicrobial activity
experiments, and 1000 mg/g. Actual silver concentrations were
593+2 and 1019+3.7 mg/g. Five strands (cut in ?2 cm long
segments) of each type of catheter were studied in this exper-
iment. The average length and weight of these strands were
71.7+0.7 cm (348+4 mg) for the catheters coated with
1000 mg/g silver, and 69.2+1.1 cm (336+5 mg) for those
Figure 2. Silver nanoparticle analysis by X-ray electron microscopy. (a) A
drop of silver nanoparticle was deposited on a carbon grid to obtain the
image (scale bar ¼ 50 nm). (b) The image was magnified and particle size
distribution was calculated using the scale bar (50 nm) from 100
measurements. (c) Higher magnification reveals the characteristic polyhedral
structure, most visible in particles 2 and 3 (scale bar ¼ 5 nm). (d) Silver
nanoparticle coating was further examined by non-disruptive electron
polycarbonate support (matrix) coated under conditions similar to those
described in the Materials and methods section (white horizontal scale bar ¼
20 nm; black vertical scale bar ¼ 20 nm). The silver coating is 80–120 nm
thick corresponding to ?4–6 layers of silver nanoparticles.
Figure 1. Silver coating of nylon catheters with silver nitrate. Coating was
performed with varying concentrations of
Materials and methods section. The y-axis represents silver deposited per
centimetre length of catheters and the x-axis represents the concentration of
silver nitrate in the coating solution. Each data point represents the mean
(+SEM) of six independent measurements. The line represents the best
(linear) fit of the data points. Error bars are smaller than the size of the
110mAgNO3as described in the
Silver nanotechnology and antimicrobial catheters
by guest on June 9, 2013
coated with 600 mg/g (mean+SEM; n ¼ 5; P ¼ not signifi-
cant). On average (10 day average), the amount of silver released
daily from catheters was 45.1+1.1 ng/cm (catheters coated
with 1000 mg/g silver) and 24.1+2.4 ng/cm (catheters coated
with 600 mg/g silver).
Daily release rates were relatively constant in both groups
although they were higher on the first days than on the last days
(data not shown). To better characterize silver release kinetics,
cumulative release data were examined (Figure 3). In 10 days,
the catheters coated with the low concentration of silver released
0.38+0.03 mg/cm of silver (14% of coated silver) and the cath-
eters coated with the higher concentration released 0.45+
1.1 mg/cm (9% of coated silver). The data show a biphasic silver
release over time, with a downward inflexion of release rates on
day 4, less marked for the catheters coated with the highest con-
centration of silver. The total amount of silver released from
these catheters in 10 days was ?20% higher than the amount of
silver released from the catheters coated with 600 mg/g silver.
On average, each animal was implanted with the equivalent of
28 cm of110mAg-coated catheters, representing 221.7+0.8 mg
of silver and 6515+24 Bq (mean+SEM; n ¼ 7). The mice
showed no sign of toxicity and looked healthy during the 10
days of the experiment. In particular, there was no sign of
inflammation or infection at the site of catheter implantation.
Further, the weight of each organ collected (heart, brain, liver,
lungs, spleen and kidneys) was similar to organ weight
measurements performed in healthy adult animals of the same
strain and sex (data not shown). On average, however, the body
weight of the animals decreased by ?8% from 36.9+2.2 g
before surgery to 34.0+0.7 g on day 10.
Urine and faeces excretion. On average, silver urine excretion
was very low (0.02 mg/day, i.e. ,0.01% of the silver present on
the catheters on day 0; Figure 4a). The total excretion of silver
in urine over the 10 day experimental period was 0.22+
0.04 mg, equivalent to 0.1% of the silver implanted at day 0. In
contrast, silver excretion in faeces was significant and varied
with time (Figure 4a). It started relatively high at 3.36+
0.44 mg on day 1 (?1.5% of implanted silver), peaked on day 2
(4.50+0.40 mg; ?2.1% of implanted silver) and then declined
in the following 8 days reaching a plateau at ?0.6–1.0 mg/day
(?0.4%) on day 6. The cumulative excretion of silver in faeces
in 10 days was 18.33+0.99 mg, equivalent to 8.3+0.4% of
the initial silver load (Figure 4b).
Organ and tissue accumulation. Silver radioactivity was deter-
mined at day 10 in several organs (heart, brain, lungs, liver,
kidney and spleen), other tissue samples (skin at the implan-
tation site, underlying muscle with ribs, control skin area, duo-
denum, caecum, femur, thigh and blood) and in the catheters.
The data are summarized in Table 2. Most of the silver remained
associated with the catheters (?84% of implanted silver). A sig-
nificant but small amount of silver was detected at the implan-
tation site (skin with panniculus carnosus and scar tissue; ?3%)
and in the muscle þ rib cage sample underlying the implantation
site (0.2%). All other organs or tissue samples including the
lungs, which under normal breathing condition, are in contact of
the rib cage had ,0.1% of the implanted silver (caecum, liver,
lungs, blood and heart) or no detectable levels of silver (control
skin sample, duodenum, spleen, kidneys, brain, femur and thigh;
data not shown). Silver recovery was high (?96% on average).
Table 1. Silver-coated catheter antimicrobial activity
Pathogen24 h48 h72 h
A count of free floating viable bacteria and the determination of viable
sessile bacteria by means of a biofilm formation assay were performed at
each time point. The results are expressed as percentage inhibition of growth
or biofilm formation versus untreated catheters. Data are presented as mean
per cent inhibition+SEM with n ¼ 12 for biofilm data and n ¼ 6 for
growth data. SEM are not shown when all data points show .99%
Figure 3. In vitro silver release kinetics from110mAg-coated catheters. Two
groups of catheters were studied: one coated with 593+2 mg of silver/g of
catheter (filled symbols) and another with 1019+3.7 mg of silver/g of
catheter (open symbols). Each data point represents the cumulative daily
average (mean+SEM; n ¼ 5) of silver released on the day indicated in
mg/cm of catheter length.
Roe et al.
by guest on June 9, 2013
The 4% unaccounted for are likely to be found at the implan-
tation site on the borders of the insertion pocket associated with
either serous liquid or scar tissue.
This report describes a method to prepare catheters coated with
silver nanoparticles and presents evidence for the catheters’ anti-
microbial properties and safety of use in animals. The method
uses silver nitrate and a mix of low-toxicity coating inducers.
The coating process is slowly reversible, yielding sustained
release of silver for at least 10 days. The released silver is active
against microorganisms most commonly found responsible for
nosocomial infections and predominantly accumulates at the site
of insertion, thus suggesting that catheters coated with this
method could provide enhanced local protection against infec-
tions with no risk of systemic toxicity.
Silver has long been known for its broad antimicrobial prop-
erties. These properties are believed to result from the disruption
of the energy metabolism and electrolyte transport systems,
which occurs when silver ions bind to bacterial sulphydryl- or
histidyl-containing proteins.17The use of silver to reduce the
risk of catheter-related infection was proposed 20 years ago by
Maki et al.,18who tested the efficacy of a biodegradable col-
lagen matrix impregnated with bactericidal silver. Since then, a
number of other methods have been developed to create cath-
eters capable of delivering silver.19These methods include elec-
tron beam assisted deposition,20,21distribution of submicron
particles of metallic silver in the polyurethane matrix of the
catheter,17placement of silver wires at the cutaneous extremity
of the catheters and release of silver by iontophoresis,22,23and
catheter impregnation with silver nanoparticles using supercriti-
cal carbon dioxide.24These methods are all relatively complex
To come up with a simpler and cheaper coating method,
silver nanoparticle production technologies were reviewed. None
of them seemed to provide a rate of nanoparticle formation and
deposition that was fast enough to be compatible with the logis-
tic of efficient industrial production. Further, many of them
utilize either starting silver salt concentrations too low (10 mM
or less) to yield potentially antimicrobial concentrations at the
surface of the catheters or agents that are toxic or at least not
biocompatible, thus precluding the use of silver nanoparticles
made by these methods because of toxicity concerns.25,26The
method described in this report is simple, uses non-toxic
chemicals and yields reproducible coating of plastic catheters
with typical nanoparticles of silver as confirmed by electron
microscopy. The amount of silver deposited is proportional to
the concentration of silver nitrate used in the coating solution
(Figure 1), at least within the range tested and thus can be
reasonably predicted. Importantly, coating takes place on both
the luminal and the external surface of the catheters, thus pro-
viding a double protection against microorganism penetration at
the implantation site. Further, the coating method does not affect
the size or the diameter of the catheters (Figure 2d) and is resist-
ant to handling. Thus, this method preserves the original qual-
ities of manufactured catheters and will likely not affect their
handling during clinical use.
Figure 4. Silver urine (open circles) and faeces (filled circles) excretion in mice implanted with110mAg-coated catheters. (a) Daily excretion. (b) Cumulative
daily excretion. Data are expressed as percentages of implanted silver (mean+SEM, n ¼ 7).
Table 2. Biodistribution of silver (mg per sample and % of
implanted silver) on day 10 (mean+SEM; n ¼ 7)
Catheters on day 10
Implantation site (skin,
panniculus carnosus, scar
Rib cage þ muscle underlying
the implantation site
All other tissues and organs: ?0.1% of implanted silver or no
Urine (cumulative 10 days)
Faeces (cumulative 10 days)
Urine and faeces data are from Figure 4.
Silver nanotechnology and antimicrobial catheters
by guest on June 9, 2013
The size distribution of the nanoparticles is relatively broad
(3–18 nm) but actually quite tight considering the simplicity of
the production process. Whether or not the distribution impacts
the microbicidal activity of the catheters is not known at this
point. One could speculate that for a given amount of coated
silver, smaller particles will yield a greater contact surface with
the liquid environment and will be more active than larger par-
ticles. Alternatively, larger particles may provide a slower but
more prolonged release of silver, a benefit for catheters destined
for longer-term implantation. The issue of size–activity relation-
ship is thus important to consider and will be the focus of future
associated with catheter use are caused by coagulase-negative
Infections caused by S. aureus, Enterococcus and E. coli are
also frequent and are often associated with increased resistance
to antibiotics. A smaller percentage of catheter-related infec-
tions are caused by other bacteria such as P. aeruginosa and
Klebsiella pneumonia, and by yeast, mainly Candida spp. Our
data show that catheters coated with silver nanoparticles are
very effective in preventing the growth of and biofilm for-
mation by most of these pathogens in vitro. Whether they will
be equally effective in vivo can only be speculated at this
time and will require further testing both in the animal model
and with controlled randomized clinical trials.
The data also show that not all microorganisms tested are
fully sensitive to silver. One could thus question the potential
advantage of catheters coated with silver nanoparticles since as
long as viable organisms remain, they will regrow as soon as
they are exposed to nutrient-containing infusion. It must be
pointedout that thesilver
described in this report are not meant to treat infections, but
rather to limit microorganism implantation, to slow their sub-
sequent growth and to retard biofilm formation. The catheters
were tested against an initial high bacterial load (1000 microor-
ganisms/mL). According to Mermel et al.,1the presence of
100 cfu/mL in blood taken from central venous catheters (500–
1000 cfu/mL in blood from peripheral intravenous catheters) is
considered positive for infection and would prompt antibiotic
treatment. In this context, silver-coated catheters should not be
considered as antimicrobial devices but as one more weapon in
the fight against infection.
The molecular mechanism underlying the antimicrobial
activity of our silver-coated catheters was not explored.
However, it is likely that growth inhibition was caused by the
silver ions (Agþ) released from the matrix of the catheter. Our
in vitro experiments indeed indicate that these catheters release
‘soluble’ silver when placed in contact with physiological saline
solution (?3 mg/day over 10 days; Figure 3). This soluble silver
may in turn have inhibited the pathogens’ respiratory enzymes
and electron transport components as proposed by others.17,27
Interestingly, the release is not constant over time. It proceeded
in two phases: a relatively fast phase during the first 4 or 5 days
and a slower phase during the next 5 days. The two-phase
release may be explained by the presence of two different pools
or layers of silver on the catheters, each having different solubi-
lity or surface density properties. Alternatively, the biphasic
release could reflect silver accessibility, the silver present on the
outer surface of the catheter being more accessible to exchange
with the solution and thus releasing faster than the silver coating
the lumen of the catheters. More extensive microscopic analysis
of the catheters after coating and at different times during
elution may be useful in the future to test these hypotheses.
Bacterial resistance to silver has been a concern with the sig-
nificant increase in the usage of silver-containing products to
manage infections.28,29Several studies suggest that it is second-
ary to a plasmid-related increase in silver binding to periplasmic
plasmid-encoded pumps.27,30The potential benefits of wide-
spread usage of silver-nanoparticle coated catheters may thus be
in part offset by the emergence of new, silver-resistant bacterial
strains. Such possibility exists with any new antimicrobial agent
but is perhaps less to fear with silver than with other agents. As
pointed out by Chopra29, target-based mutation to silver resist-
ance is unlikely because of the multiplicity of intracellular
targets of silver ions and may explain in part the limited number
of reports of silver resistance in bacteria published so far.
Catheters coated with silver nanoparticles could be used to
perfuse various types of solutions. Some of these solutions may
be poorly compatible with the silver coating and either damage
it or reduce its antimicrobial activity. Laboratory experiments
have shown that the coating is stable under pH conditions
ranging from 2 to 12, hence the use of 2% HNO3as a rinsing
agent in the final steps of the coating process. Thus, the pH of
commonly used injectable solutions is unlikely to damage the
coating of the catheters. Cysteine-, SH- or sulphide-containing
compounds as may be found in albumin or amino acid solutions
may bind silver ions and enhance their elution from the coating
of the catheters. Chloride ions and phosphate ions, as found in
many injectable solutions may also react with silver ions but
decrease their release from the coating by forming an insoluble
layer of AgCl or Ag3PO4at the surface of the nanoparticles.
Compatibility issues cannot be fully predicted at this time and
will need to be examined on a case by case basis. It must be
pointed out though that if the dissociation constants of AgCl and
Ag3PO4 are low (1.6 ? 10210and 1.6 ? 1025M at 258C,
respectively), they are nonetheless high enough to allow some of
the silver to be released as Agþ. Further, as shown by the data
presented in Table 1, significant antimicrobial activity of the
silver-coated catheters was observed in a medium containing
both a protein digest (peptone) and sodium chloride. Thus,
chemical interactions between silver-coated catheters and per-
fusion solutions may occur and change the kinetics of the
release of silver ions from the catheters but are unlikely to sig-
nificantly reduce overall Agþ
The compatibility between injectable solutions and the silver-
coated catheters may not be so much of a concern. Parenteral
solutions are sterile and catheter-related infections are primarily
caused by microorganisms invading the site of insertion of the
catheter and the tissues in contact with the outside of the cath-
eter. Of potentially greater concern is the possibility of systemic
toxicity from the silver eluted from the catheter. Although silver
is a low toxicity metal,31–33this issue was addressed by evaluat-
ing the biodistribution of silver released from silver-coated cath-
eters in the mouse.
The animals lost on average 8% of their body weight
throughout the 10 day experimental period. Such a decrease is
likely secondary to the change in housing conditions (from shoe
box before surgery to individual metabolic cage after surgery)
but could be attributable to silver toxicity. The data however
availability for bactericidal
Roe et al.
by guest on June 9, 2013
show that in 10 days the catheters released ?16% of their
coating. The majority of the released silver is excreted in the
faeces (?50%) and a significant amount remains associated
with the tissues in direct contact with the catheters (?20%). No
significant accumulation of silver was observed in the major
organs examined. These data are in keeping with previous
reports in rodents and other species.34,35The amount of silver
released from a 28 cm long catheter strand over the 10 day
period is equal to ?35 mg. If all this silver accumulated in
tissues, it would amount to a dose of ?1 mg/kg mouse body
weight and 0.5 mg/kg human body weight (for an adult of
70 kg). With a LD50 in the g/kg body weight range (in the
mouse), such exposure to silver rules out the possibility that the
animal weight loss was caused by the silver released by the cath-
eters and suggests that the catheters coated with the silver nano-
particle technology will not cause any systemic toxicity in
humans. The biodistribution data further indicate that there is a
4000-fold silver gradient over a distance of ?3–10 mm (from
the catheter to the lungs and heart). This suggests a very limited
diffusion of released silver through tissues and a very limited
risk for subcutaneous catheters coated with silver to release
toxic concentrations of silver to underlying tissues and organs in
The excretion of silver in faeces, the major route of silver
elimination in vivo, follows a biphasic pattern, with a rapid
excretion rate for the first 3–4 days followed by a slower
excretion rate from day 4 to day 10 (Figure 4). This mirrors the
silver elution patterns observed in vitro, suggesting that silver
release from implanted catheters follows the same kinetics in
vivo and in vitro. Thus, the in vitro elution test might be an effi-
cient and economic alternative to animal testing when it comes
to predict the biological properties of new catheter formulations
or other medical devices treated with our method.
In conclusion, a method to coat plastic catheters with bio-
active silver nanoparticles was developed. These catheters are
non-toxic devices capable of targeted and sustained release of
bactericidal silver at the implantation site and may prove useful
in preventing infectious complications in patients with indwel-
This work was supported in part by a grant from the National
Institutes of Health (R43 AI061894) awarded to AcryMed, Inc.,
and by AcryMed, Inc. R&D.
Dr N. B.-S.: none to declare. Dr J.-B. R. was the recipient of a
NIH subcontract from AcryMed, Inc. (AcryMed, Inc./Dr
B. G. primary recipient). Drs B. G., B. K and D. R. own stocks
or shares in a company that could be financially affected by the
conclusions of this article. Dr B. G. is Chief Technical Officer
and Chairman of AcryMed, Inc.
1. Mermel LA, Farr BM, Sheretz RJ et al. Guidelines for the man-
agement of intravascular catheter-related infections. Clin Infect Dis
2001; 32: 1249–72.
2. Rosenthal K. Guarding against vascular site infection: arm your-
self with the latest knowledge on equipment and technique to protect
patients from catheter-related bloodstream infections. Nurs Manage
2006; 37: 54–66.
3. Randolph AG, Brun-Buisson C, Goldmann D. Identification of
central venous catheter-related infections in infants and children.
Pediatr Crit Care Med 2005; 6 Suppl 3: 19–24.
4. Kline A. Pediatric catheter-related bloodstream infections. Latest
strategies to decrease risk. AACN Clin Issues 2005; 16: 185–98.
5. CDC NNIS System. National nosocomial infections surveillance
(NNIS) system report, data summary from January 1992 through June
2004, issued October 2004. Am J Infect Control 2004; 32: 470–85.
6. Johnson JR, Kuskowski MA, Wilt TJ. Systematic review: antimi-
crobial urinary catheters to prevent catheter-associated urinary tract
infection in hospitalized patients. Ann Intern Med 2006; 144: 116–26.
7. O’Grady NP, Alexander M, Dellinger EP et al. Guidelines for the
prevention of intravascular catheter-related infections. Pediatrics 2002;
8. Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a
common cause of persistent infections. Science 1999; 284: 1318–22.
9. Hanna H, Raad I. New approaches for prevention of intravascu-
lar catheter-related infections. Infect Med 2001; 18: 38–48.
10. Walder B, Pittet D, Tramer MR. Prevention of bloodstream infec-
tions with central venous catheters treated with ant-infective agents
depends on catheter type and insertion time: evidence from a
meta-analysis. Infect Control Hosp Epidemiol 2002; 23: 748–56.
11. Hanna HA, Raad II, Hackett B et al. Antibiotic-impregnated cath-
multidrug-resistant bacteremias in critically ill patients. Chest 2003;
12. Chaiban G, Hanna H, Dvorak T et al. A rapid method of impreg-
nating endotracheal tibes and urinary catheters with gendine: a novel
antiseptic agent. J Antimicrob Chemother 2005; 55: 51–6.
13. Hawser SP, Norris H, Jessup CJ et al. Comparison of a
H-tetrazolium hydroxide (XTT) colorimetric method with the standar-
dized National Committee for Clinical Laboratory Standards method of
testing clinical yeast isolates for susceptibility to antifungal agents.
J Clin Microbiol 1998; 36: 1450–2.
14. Tunney MM, Ramage G, Field TR et al. Rapid colorimetric
assay for antimicrobial susceptibility testing of Pseudomonas aerugi-
nosa. Antimicrob Agents Chemother 2004; 48: 1879–81.
15. Zheng J-G. Silver nanoprisms and nanotetrahedra investigated
by transmission electron microscopy. Microsc Microanal 2003; 9:
16. Yang Z, Chang H-T. Anisotropic syntheses of boat-shaped core-
shell Au-Ag nanocrystals and nanowires. Nanotechnology 2006; 17:
17. Guggenbichler JP, Boswald M, Lugauer S et al. A new technol-
ogy of microdispersed silver in polyurethane induces antimicrobial
activity in central venous catheters. Infection 1999; 22: S16–23.
18. Maki DG, Cobb L, Garman JK et al. An attachable silver impreg-
nated cuff for prevention of infection with central venous catheters: a
prospective, randomized multicenter trial. Am J Med 1988; 85: 307–14.
19. Sticler DJ. Biomaterials to prevent nosocomial infections: is
silver the gold standard? Curr Opin Infect Dis 2000; 13: 389–93.
20. Sioshansi P. New processes for surface treatment of catheters.
Artif Organs 1994; 18: 266–71.
Silver nanotechnology and antimicrobial catheters
by guest on June 9, 2013
21. Trerotola SO, Johnson MS, Shah H et al. Tunneled hemodialy- Download full-text
sis catheters: use of a catheter for prevention of infection—a random-
ized study. Radiology 1998; 207: 491–6.
22. Raad I, Hachem R, Zermeno A et al. In vitro antimicrobial effi-
cacy of silver iontophoretic catheter. Biomaterials 1996; 17: 1055–9.
23. Raad I, Hachem R, Zermeno A et al. Silver iontophoretic cath-
eter: a prototype of a long-term antiinfective vascular access device.
J Infect Dis 1996; 173: 495–8.
24. Furno F, Morley KS, Wong B et al. Silver nanoparticles and poly-
meric medical devices: a new approach to prevention of infection?
J Antimicrob Chemother 2004; 54: 1019–24.
25. Nickel U, Castell AZ, Poppl K et al. A silver colloid produced by
surface-enhanced Raman spectroscopy. Langmuir 2000; 16: 9087–91.
26. Green M, Allsop N, Wakefield G et al. Trialkylphosphine oxide/
amine stabilized silver nanocrystals—the importance of steric factors
and Lewis basicity in capping agents. J Mater Chem 2002; 12:
27. Li X-Z, Nikaido H, Williams K. Silver-resistant mutants of
Escherichia coli display active efflux of Ag and are deficient in porins.
J Bacteriol 1997; 179: 6127–32.
28. Percival SL, Bowler PG, Russell D. Bacterial resistance to silver
in wound care. J Hosp Infect 2005; 60: 1–7.
29. Chopra I. The increasing use of silver-based products as antimi-
crobial agents: a useful development or a cause of concern?
J Antimicrob Chemother 2007; 59: 587–90.
30. Silver S. Bacterial silver resistance: molecular biology and uses
and misuses of silver compounds. FEMS Microbiol Rev 2003; 27:
31. Armitage SA, White MA, Wilson HK. The determination of silver
in whole blood and its application to biological monitoring of
32. Saber H, Anner RM, Anner BM. Cysteine protects Naþ/
Kþ-ATPase and isolated human lymphocytes from silver toxicity.
Biochem Biophys Res Commun 1992; 189: 1444–9.
33. Wright JB, Lam K, Buret AG et al. Early healing events in a
porcine model of contaminated wounds: effects of nanocrystalline
silver on matrix metalloproteinases, cell apoptosis, and healing. Wound
Rep Reg 2002; 10: 141–51.
34. Furchner JE, Richmond CR, Drake GA. Retention of silver-110m
in the mouse, rat, monkey and dog. Health Phys 1987; 15: 505–14.
35. Gammill JC, Wheeler B, Carothers EL et al. Distribution of radio-
active silver colloids in tissues of rodents following injection by various
routes. Proc Soc Exp Bio Med 1950; 74: 691–5.
Am Occup Hyg1996;40:
Roe et al.
by guest on June 9, 2013