Acta Biomaterialia 6 (2010) 3791–3797

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Acta Biomaterialia

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Biofilm formation on bone grafts and bone graft substitutes: Comparisonof different materials by a standard in vitro test and microcalorimetry

Martin Clauss a,b,c,*, Andrej Trampuz c,d, Olivier Borens d,e, Marc Bohner b, Thomas Ilchmann a

a Department of Orthopedic Surgery, Kantonsspital Liestal, Liestal, Switzerlandb RMS Foundation, Bettlach, Switzerlandc Infectious Diseases Service, Department of Internal Medicine, University Hospital Lausanne (CHUV), Lausanne, Switzerlandd Septic Surgical Unit, Department of Surgery, University Hospital Lausanne (CHUV), Lausanne, Switzerlande Department of Orthopedics and Traumatology, University Hospital Lausanne (CHUV), Lausanne, Switzerland

a r t i c l e i n f o a b s t r a c t

Article history:Received 3 December 2009Received in revised form 14 February 2010Accepted 5 March 2010Available online 11 March 2010

Keywords:BiofilmBone graftb-TCPInfectionMicrocalorimetry

1742-7061/$ - see front matter � 2010 Acta Materialdoi:10.1016/j.actbio.2010.03.011

* Corresponding author. Address: Kantonsspital Liedic Surgery, Rheinstrasse 26, CH-4410 Liestal, Switzefax: +41 61 925 2808.

E-mail address: martin.clauss@ksli.ch (M. Clauss).

We analyzed the initial adhesion and biofilm formation of Staphylococcus aureus (ATCC 29213) and S. epi-dermidis RP62A (ATCC 35984) on various bone grafts and bone graft substitutes under standardizedin vitro conditions. In parallel, microcalorimetry was evaluated as a real-time microbiological assay inthe investigation of biofilm formation and material science research. The materials b-tricalcium phos-phate (b-TCP), processed human spongiosa (Tutoplast™) and poly(methyl methacrylate) (PMMA) wereinvestigated and compared with polyethylene (PE). Bacterial counts (log10 cfu per sample) were higheston b-TCP (S. aureus 7.67 ± 0.17; S. epidermidis 8.14 ± 0.05) while bacterial density (log10 cfu per surface)was highest on PMMA (S. aureus 6.12 ± 0.2, S. epidermidis 7.65 ± 0.13). Detection time for S. aureus bio-films was shorter for the porous materials (b-TCP and processed human spongiosa, p < 0.001) comparedto the smooth materials (PMMA and PE), with no differences between b-TCP and processed humanspongiosa (p > 0.05) or PMMA and PE (p > 0.05). In contrast, for S. epidermidis biofilms the detection timewas different (p < 0.001) between all materials except between processed human spongiosa and PE(p > 0.05). The quantitative analysis by quantitative culture after washing and sonication of the materialdemonstrated the importance of monitoring factors like specific surface or porosity of the test materials.Isothermal microcalorimetry proved to be a suitable tool for an accurate, non-invasive and real-timemicrobiological assay, allowing the detection of bacterial biomass without removing the biofilm fromthe surface.

� 2010 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

1. Introduction metal meshes). Bone grafts become incorporated by the host bone

Bone transplantation is the most commonly performed trans-plantation, performed about 10 times more often than any othersolid organ transplantation [1]. For filling bone defects, autologousor allogeneic bone grafts or bone graft substitutes are used. Cur-rently, mainly cancellous autologous bone transplants are im-planted, typically harvested from the iliac crest or the distalradius and transferred as fresh autologous grafts during the sameoperation [1,2]. For allogeneic bone grafts, femoral heads are har-vested during hip replacement surgery and stored as deep frozengrafts in a bone bank. Filling bone defects with artificial materialsincludes bone grafts (such as b-tricalcium phosphate (b-TCP),hydroxyapatite (HA) or processed spongiosa) and bone graft sub-stitutes (such as poly(methyl methacrylate) (PMMA) or tantalum

ia Inc. Published by Elsevier Ltd. A

stal, Department of Orthope-rland. Tel.: +41 61 925 2525;

and are substituted either completely (b-TCP, processed spongiosa)or partially (HA), while bone graft substitutes are not biodegrad-able and thus remain incorporated in the bone unchanged.

Infection of the surgical site is a recognized complication in thevicinity of a foreign body. Despite antimicrobial prophylaxis andthe use of laminar-flow ventilation in operating rooms, surgicalsite infections cannot be prevented completely [3]. Infection ratesrange from 0.7% to 4.2% in elective orthopedic surgery [4] and canreach up to 30% following third-degree open fractures [5]. Sincebone grafts and bone graft substitutes are being used increasinglyin orthopedic surgery, we will face an increasing number of in-fected bone grafts and bone graft substitutes in the future. Autoge-neous and allogeneic bone grafts are temporary or permanentforeign bodies that are devoid of vascularization and are thereforeareas of reduced resistance against the bacteria [6] that are eitherintroduced during surgery or reach the implant after surgery eithervia the skin (per continuitatem) or via the blood. The most com-mon infecting microorganisms (70–90%) are staphylococci [7–9],which typically grow attached on the surface as a specific structure

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3792 M. Clauss et al. / Acta Biomaterialia 6 (2010) 3791–3797

known as a biofilm [7,10]. The treatment of infections caused bybiofilms is more difficult than of bacteria growing in free-living(planktonic) form. The eradication of infection is often only possi-ble by removal of the implant and long-term antimicrobial treat-ment [11]. The ‘‘race to the surface” [8] among bacteria and hostcells is influenced by various factors, which have been much stud-ied in the metal implants used as fracture-fixation devices or jointprostheses. For these implants, the first step of infection (bacterialadhesion) and the later developments (biofilm formation) are af-fected by the material type and surface properties [8,12–17]. Therisk of infection associated with bone grafts or bone graft substi-tute in vivo is unknown.

In this study, we evaluated the initial adhesion and biofilm for-mation under standard in vitro conditions using two laboratorystrains of bacteria commonly causing surgical site bone infection,namely Staphylococcus aureus (ATCC 29213) and S. epidermidisRP62A (ATCC 35984). Various bone grafts and bone graft substi-tutes were chosen to obtain a representative selection of clinicallyimportant biomedical materials.

In parallel, we investigated whether microcalorimetry is a suit-able tool with which to investigate biofilm formation on the sur-face of bone grafts and bone graft substitutes. The heat flow isproportional to the quantity of bacteria, and can be used for quan-titative assessment [18–20]. We hypothesized that microcalorime-try can be used as an accurate, real-time microbiological assay inthe investigation of biofilm formation and material science re-search. It has the advantage of being a non-invasive approach,allowing the detection of bacterial biomass without removing thebiofilm from the surface [21,22]. In order to validate the microca-lorimetric method, experiments were performed in parallel withconventional quantitative cultures of the sonication fluid obtainedafter washing and sonication of the material [23].

2. Materials and methods

The following bone grafts and bone graft substitutes were usedas test samples:

� Bone grafts. b-TCP (RMS Foundation, Bettlach, Switzerland), pro-duced by the so-called ‘‘calcium phosphate emulsions” method(comparable to chronOS™, Synthes Biomaterials, Bettlach, Swit-zerland), with a porosity ranging from 72% to 75%, a macroporediameter close to 0.4 mm and some degree of interconnectivity[24,25]. b-TCP is a ceramic, so it is by definition brittle and itsshear and tensile properties are poor [26,27]. Upon implanta-tion, b-TCP is replaced by mature bone within several monthsto years [27–29].� Processed human spongiosa (Tutoplast™, Novomedics GmbH,

Zürich, Switzerland), a human graft material after a multi-stepchemical treatment (osmolysis, NaOH, H2O2, acetone) [27].Samples show an interconnecting porosity with a pore size ofseveral 100 lm. The material has a mineral phase of about65 wt.% (49 wt.% carbonated hydroxyapatite). The remaining33 wt.% is due to water and persistent bone matrix [27]. Itsintrinsic mechanical stability is moderate [27]; nevertheless,in the case of mechanical loading, the graft must be augmented.� Bone graft substitute. Bone cement blocks (Biomet Bone Cement

R™, Biomet Orthopedics GmbH, Ried, Switzerland): PMMA is ahydrophilic acrylic resin, well established for filling bonydefects [26], which was hand-mixed in a bowl without vacuumapplication. The outer form can be molded in the liquid phaseand, after thermal setting, forms a solid block, providing excel-lent mechanical stability. The material has no osteoinductive orosteoconductive capacity and remains inert with the surround-ing host tissue, covered in a fibrous tissue envelope [26].

Biofilm formation on ultrahigh-molecular-weight polyethylene(PE; RMS Foundation, Bettlach, Switzerland) has been extensivelystudied previously [30,31]. In this study, we included PE as a con-trol material. PE blocks were machined out of a single solid bar,washed in ethanol and sterilized by gamma irradiation.

2.1. Manufacture and characterization of test materials

All test samples were manufactured as cylinders (6.5 mm diam-eter � 10 mm height) ready for ‘‘off-the-shelf” use. Exact samplesizes were determined by measuring the diameter and heightusing a caliper instrument (precision 0.01 mm) and block volumeswere calculated. Sample weight was determined with an electronicprecision balance (Mettler AE 260 Delta range, precision 0.0001 g)and sample porosity was calculated. For the porous blocks (b-TCP,processed human spongiosa) the material surface was measuredby nitrogen gas adsorption (BET) [32]. The surface of the solidblocks (PE and PMMA) was approximated to be equal to the calcu-lated cylinder surface. Sample surfaces were additionally analyzedby light microscopy and scanning electron microscopy (SEM).

2.2. Bacterial strains

Two representative staphylococcal strains were used [30,33]: S.aureus (ATCC 29213) is a Gram-positive, coagulase-positive, meth-icillin-susceptible, biofilm-forming strain [33]; S. epidermidisRP62A (ATCC 35984) is a Gram-positive, coagulase-negative, ica-positive [34], biofilm-forming strain with a glycocalyx capsule[35]. The strains were stored at �70 �C by using a cryovial beadpreservation system (Microbank, Pro-Lab Diagnostics, RichmondHill, Ontario, Canada). For preparation of the inoculum, a singlebead was freshly grown on sheep blood agar overnight. Bacterialinocula were prepared from discrete colonies resuspended in 1%phosphate-buffered saline (PBS) to a McFarland turbidity of 0.5,representing a bacterial concentration of 9.7 ± 0.8 � 107 colony-forming units (cfu) ml�1 for S. aureus and 1.3 ± 0.3 � 107 cfu ml�1

for S. epidermidis. This stock solution was diluted 1:100 for furtherexperiments.

2.3. Biofilm formation

Fifty-milliliter Falcon tubes were pre-filled with 2700 ll trypticsoy broth (TSB; Beton Dickinson AG, Basel, Switzerland). Six repli-cates of each test material were inserted onto the surface of thebroth with a sterile forceps to allow a homogeneous soaking ofthe porous materials [44] over a 30 min period. In a final step300 ll of diluted bacterial stock solution was added. Samples wereincubated at 37 �C with ambient air for 24 h without shaking.

2.4. Quantification of adherent biofilm

2.4.1. By sonicationAfter 24 h, the test samples were transferred to a new 50 ml Fal-

con tube with a sterile forceps and gently washed three times with1% PBS. The samples were then transferred to 3 ml of fresh PBS,vortexed (Vortex Genie 2, Scientific Industries, Bohemia NY, USA)for 10 s with maximum power, sonicated (BactoSonic™, Bandelinelectronic, Berlin, Germany) at 40 kHz for 5 min and vortexedagain for 10 s [23]. Sonication fluid was transferred to a 10 ml Fal-con tube, plated within 30 min in aliquots of 100 ll onto aerobicblood agar with 5% sheep blood and incubated at 37 �C with ambi-ent air for 24 h. Bacterial counts were enumerated and expressedas cfu ml�1. The remaining bone grafts and bone graft substituteswere harvested for further investigation and transferred to themicrocalorimeter, where they were processed immediately.

M. Clauss et al. / Acta Biomaterialia 6 (2010) 3791–3797 3793

2.4.2. By microcalorimetryAfter sonication the test samples were transferred to sterile

4 ml calorimeter ampoules pre-filled with 1 ml TSB, closed with arubber cap and sealed by manual crimping. Ampoules weresequentially introduced into the calorimetry instrument and re-mained for 15 min in the thermal equilibration position beforebeing lowered into the measurement position. A 48-channel batchcalorimeter (thermal activity monitor, model 3102 TAM III; TAInstruments, New Castle, DE, USA) was used to measure the heatflow at 37 �C controlled to within 0.0001 �C and had a sensitivityof 0.25 lW. Heat flow was measured after the signal stabilitywas achieved after 45 min at 10 s intervals during 24 h. The batchmode gives the sum signal of both surface-associated as well asplanktonic growing cells.

Heat was measured continuously and expressed as heat flowover time (in microwatts). Calorimetric time to detection (TTD)was defined as the time from insertion of the ampoule into the cal-orimeter until the exponentially rising heat flow signal exceeded75 lW for S. epidermidis and 100 lW for S. aureus. TTD indirectlyquantifies the amount of bacteria, with a shorter TTD representinga larger amount of bacteria. The peak heat flow (PHF) was definedas the maximum heat flow during the experiment, and the time toreach the peak heat flow (ttPHF) was recorded. PHF is correlatedwith two factors: (i) linearly with the volume of growth mediumadded to the ampoules; and (ii) in case of equal volumes of growthmedium individual to the strain (unpublished data). ttPHF, likeTTD, represents a measure of the bacterial load. The total producedheat (TPH; in joules) was determined by integrating the area belowthe heat flow–time curve. Data analysis was performed using thecalorimeter manufacturer’s software (TAM Assistant; TA Instru-ments) and Prism 5.0 (GraphPad Software, La Jolla, CA).

2.5. Statistical calculations

All experiments were performed in triplicate to calculate themean and standard deviation. To equalize variances in bacterialcounts, data are presented as log10. Additionally the numbers ofcfu are normalized to the sample surface by the equation: cfuper surface = cfu per sample � average surface sample/average sur-

Fig. 1. Material characteristics. Values present mean values ± SD of 10 repeated measuremmeasurements of nitrogen gas adsorption.

face PE. For statistical analysis, one-way analysis of variance withBonferroni’s multiple comparison test was used. A p-value of<0.05 (two-sided) was considered statistically significant.

3. Results

3.1. Material characteristics

The sample size parameters were consistent for each material,but the examined materials significantly differed in diameter,height, volume, porosity and surface area (Fig. 1). However,whereas the differences in diameter, height and volume were onlyin the 1–2% range, the SSA values varied over three orders of mag-nitude, ranging from a few mm2 for PE and PMMA to 1000 mm2 forb-TCP and processed human spongiosa. The latter difference re-sults from the presence of micropores in b-TCP and processed hu-man spongiosa, as revealed by light microscopy (Fig. 2). Thesephotos also show that the porosity of b-TCP is more homogeneousthan that of processed human spongiosa, probably because b-TCPis a synthetic material whereas processed human spongiosa is de-rived from natural bone. The PMMA surface was smooth with somepores (Fig. 2g) and areas of imperfect homogenization betweenpores (Fig. 2k) due to hand mixing. Finally, the PE surfaces pre-sented typical signs of machining but no pores or defects.

3.2. Biofilm formation

Both test strains formed significant biofilm on test materials, asconfirmed by quantitative culture after washing and sonication ofthe material. In the inoculation culture used for induction of bio-film formation, the bacterial concentration after 24 h incubationin TSB was 1.25 ± 0.2 � 1010 cfu ml�1 with S. aureus and7.9 ± 0.1 � 108 cfu ml�1 with S. epidermidis.

3.3. Quantification of adherent biofilm

3.3.1. By sonicationFor S. aureus, bacterial counts (log10 cfu per sample) were higher

(p < 0.001) from porous (b-TCP, 7.67 ± 0.17; processed human

ents (diameter, height and weight) or three repeated measurements (surface). BET:

3794 M. Clauss et al. / Acta Biomaterialia 6 (2010) 3791–3797

spongiosa, 7.65 ± 0.15) than from solid materials (PMMA,6.12 ± 0.18; PE, 5.17 ± 0.22). No difference in bacterial density(log10 cfu per surface) was observed between b-TCP, processed hu-man spongiosa and PE (p > 0.05), but a significant higher bacterialdensity was detected on PMMA (p < 0.001) (Fig. 3a). Staphylococcus

Fig. 3. Bacterial counts in sonication fluid after removal from different test sample

Fig. 2. Surface configuration (porosity and roughness) of the tested materials. (First row1 mm. (Middle and bottom rows) SEM analysis. The bars represent 200 lm (middle row

epidermidis showed significantly higher bacterial counts (log10 cfuper sample) on b-TCP (8.14 ± 0.05, p < 0.001) than on solid samples(PMMA 7.71 ± 0.09, PE 7.52 ± 0.09), but no differences were ob-served between both solid samples (PMMA and PE). The bacterialdensity (log10 cfu per surface) was highest on PMMA

s. (a) S. aureus; (b) S. epidermidis. Data represent mean ± SD of two triplicates.

) The front side of the samples as shown using light microscopy. The bar represents) and 50 lm (bottom row), respectively.

Fig. 4. Microcalorimetric results for (a) S. aureus and (b) S. epidermidis. Datarepresent triplicates for each test material. Dotted line indicates TTD for S. aureus(100 lW) and S. epidermidis (75 lW). The negative values at the beginning of theexperiment are due to the setting (signal stability achieved after 45 min) of themicrocalorimeter, not the experimental conditions.

M. Clauss et al. / Acta Biomaterialia 6 (2010) 3791–3797 3795

(7.65 ± 0.13) and was increased by about one-third compared to S.aureus (6.12 ± 0.2, p < 0.001). The bacterial density (log10 cfu persurface) on the PE samples was more than 100 times higher forthe S. epidermidis than for the S. aureus (7.47 ± 0.09 vs.5.18 ± 0.24) (Fig. 3b).

3.3.2. By microcalorimetryNo detectable heat flow was observed in the ampoules filled

with TSB alone or in those with TSB and each of the four test sam-ples (negative control). The shapes of the heat flow curves werecharacteristic for the individual bacterial strains and were notinfluenced by the test materials. The calorimetric time to detectionwas shortest for b-TCP for both strains, and was shorter for S. aur-eus than for S. epidermidis on all examined materials (Table 1).

For S. aureus, TTD was similar for both porous materials (b-TCPand processed human spongiosa; p > 0.05) and for both smoothmaterials (PMMA and PE; p > 0.05), but was shorter for the porousthan for the smooth materials (p < 0.001). A shorter TTD indicates ahigher number of bacteria. In contrast, for S. epidermidis the time todetection was different (p < 0.001) between all materials despiteprocessed human spongiosa and PE (p > 0.05). PHF was about twiceas high for S. aureus, (207 ± 16 lW) as for S. epidermidis(102 ± 5 lW) independent from the test material. ttPHF was short-er for S. aureus than for S. epidermidis with corresponding materialsexcept with PE (Fig. 4a and b and Table 1). No difference in ttPHFwithin either the two porous or the two smooth materials was ob-served, though there was a difference between porous and smoothmaterials (p < 0.001) for S. aureus. TPH was highest on the PE con-trol samples for S. aureus 5.76 ± 0.56 J (p < 0.001) and S. epidermidis4.17 ± 0.27 J (p < 0.001), but with no significant difference betweenthe other tree tested materials (p > 0.05, Fig. 4 and Table 1).

4. Discussion

Extensive research has been performed in recent years to deter-mine the propensity of medical devices to sustain biofilm forma-tion by staphylococci [17]. Several factors of the material, such assurface hydrophobicity [36], surface charge [15], surface roughness[16] and surface chemistry [35,37,38], have been shown to influ-ence biofilm formation. Therefore, surface modifications may influ-ence bacterial adhesion and the related risk of infection. In general,porous materials have a higher rate of infection than dense mate-rials [30]. In addition, S. aureus preferably adheres to metals, whileS. epidermidis adheres better to polymers [8,15]. In this study, testmaterials differing in structural, physical and chemical propertieswere selected to represent bone grafts and bone graft substitutescurrently available on the market. All materials were quoted bythe manufacturers as ‘‘off-the-shelf” preparations to simulate anin vitro situation as close as possible to the clinical setting.

Table 1Biofilm formation of S. aureus and S. epidermidis on different test samples evaluatedby microcalorimetry.

Strain Material TTD (h) PHF (lW) ttPHF (h) TPH (J)

S. aureus b-TCP 1.4 ± 0.1 210.8 ± 11.7 3.0 ± 0.2 4.3 ± 0.3Tutoplast 1.5 ± 0.4 208.3 ± 13.3 3.0 ± 0.3 4.2 ± 0.7PMMA 5.6 ± 0.2 190.0 ± 8.4 7.1 ± 0.3 4.2 ± 0.3PE 5.6 ± 0.4 218.7 ± 13.1 7.1 ± 0.5 5.8 ± 0.6

S. epidermidis b-TCP 2.0 ± 0.1 108.5 ± 1.4 3.9 ± 0.1 3.3 ± 0.1Tutoplast 4.9 ± 0.6 98.9 ± 4.0 7.1 ± 0.5 3.9 ± 0.2PMMA 10.1 ± 0.7 102.4 ± 2.8 12.2 ± 0.6 3.6 ± 0.2PE 4.0 ± 0.3 98.2 ± 2.8 6.5 ± 0.6 4.2 ± 0.3

Data are presented as means ± SD. TTD was measured at 100 lW for S. aureus and75 lW for S. epidermidis. PHF reflects the highest point of the heat flow curve andttPHF the time period until PHF was reached. TPH represents the total amount ofenergy produced during the 24 h interval.

We used staphylococcal strains that have been previouslyshown to be good biofilm formers, producing a mature biofilm after24 h for S. aureus [39] and S. epidermidis [34,40,41], when incubatedin TSB [30,35,39–41]. A number of publications have establishedthat the two standard strains used for this study are biofilm form-ers. Choosing standard (ATCC) strains rather than clinical strains al-lows other researchers to repeat the studies and compare theirresults. The choice of growth medium (human serum, TSB, CaMHB,PBS) was shown to influence biofilm formation considerably, as itchanges the initial bacterial adsorption on the surface [31,35]. Hu-man serum decreased the initial bacterial adhesion up to 6 h. Thiswas followed by increased biofilm formation after 12–24 h for theS. epidermidis strain RP62A [35]. We selected TSB for the in vitroevaluation of biofilm formation since this is the most commonlyused medium in previous studies [30,35,39]. Gentle shaking wasemployed to improve biofilm formation, as done previously [39],except for fragile samples (b-TCP and processed human spongiosa),where shaking might have modified the surface, so static growthconditions were chosen. It is essential that the removal of plank-tonic bacteria by only the biofilm is monitored. Washing test sam-ples with PBS [41], saline [30] or deionized water [36] is anestablished method to remove planktonic bacteria, leaving onlythe attached biofilms on the surface remaining. This process couldbe critical for porous materials like b-TCP and processed humanspongiosa because it is possible that not all the planktonic bacteriawill be washed out of the micropores, resulting in an overestima-tion of the true amount of biofilm. Therefore, the washing proce-dure was uniformly performed three times for all samples. Afterwashing, sonication was used to dislodge biofilms from devices[39], as adapted from a recently published protocol [23].

Our results demonstrate that bacterial counts (cfu per sample)removed after sonication are about 10-fold (S. epidermidis) or100-fold (S. aureus) higher on porous samples (b-TCP, processedhuman spongiosa) compared to smooth samples (PMMA and PE).The bacterial counts obtained from sonication fluid showed low

3796 M. Clauss et al. / Acta Biomaterialia 6 (2010) 3791–3797

variability within the replicates for S. aureus and S. epidermidis,demonstrating a reliable model for quantitation of biofilm forma-tion on three-dimensional test samples, independent on porosity.The bacterial density in the biofilm was higher with S. epidermidisthan with S. aureus (except for processed human spongiosa), de-spite a higher bacterial concentration of S. aureus in the inoculationculture. This might represent the evolutionary adaptation of thelow-virulence S. epidermidis to form a dense biofilm, while themore virulent S. aureus has a propensity to grow preferably in aplanktonic state.

Bacterial density (cfu per surface) was higher on the sampleswith smooth surfaces (PMMA and PE) for both strains comparedto the porous/rough samples, which is in contrast to findings onsteel and titanium alloys [16]. Biofilm formation is a multifactorialprocess and differences in surface roughness and the resulting in-crease in surface area may be more important for solid samplesthan for porous samples. We also found a difference in bacterialdensity on PMMA samples compared to PE for S. aureus but notfor S. epidermidis. Oga et al. [15] found higher bacterial quantitieson PMMA compared to PE when using another S. epidermidis strain(E-46) in a dynamic flow chamber system filled with PBS to pro-duce the biofilm. The differences in the conditions of biofilm for-mation (flow chamber/PBS vs. static conditions/TSB) are apossible explanation for these different findings, and the two dif-ferent S. epidermidis strains (E-46 vs. RP62A) used might also havecontributed.

Transfer of the samples to calorimetry ampoules after sonicationmakes a direct correlation with the bacterial counts more difficult assome of the biofilm is dislodged and the number of adherent bacteriais reduced by this procedure. Microcalorimetry, in addition to beingmore sensitive and easier to perform, correlated well with the soni-cation results (Figs. 3 and 4). As we were interested in quantitativelymonitoring the mature part of a 24 h biofilm on the surface of variousirregularly shaped bone grafts and bone graft substitutes and evalu-ating calorimetry as a potential tool, we believe that removingplanktonic bacteria by sonication and perhaps also an immature partof the biofilm is more important than to comparing the differencesbetween sonication and standard plating. Furthermore the experi-mental setting shows that the percentage of the biofilm dislodgeddiffers with the strains and materials used.

For planktonic bacteria, calorimetry has been widely used as a‘‘non-invasive” [42] monitoring instrument due to its high sensitiv-ity, reproducibility and simplicity [19]. There are various othermethods for quantitative/qualitative evaluation of biofilm forma-tion, such as ‘‘live–dead staining” [43], confocal laser scanningmicroscopy (CLSM) [34,43], fluorescence microscopy [16,36], elec-tron microscopy (transmission electron microscopy/SEM)[15,16,41] or atomic force microscopy (AFM) [41]. All methods butcalorimetry need a special pre-treatment, like staining (live-deadstaining, CLSM) or carbon-sputtering (reflection electron micros-copy/SEM), which an further hinder biofilm investigation afterquantification [43] or make it impossible to assess on rough orthree-dimensional porous structures (AFM; unpublished data San-dor Kasas, EPFL, Lausanne, Switzerland). Furthermore, the area/vol-ume that can be monitored by other methods is limited, and theresults might be biased from choosing a specific area while microcal-orimetry monitors the whole device. Quantitative microcalorimetricmonitoring of living bacteria growing in a biofilm is a reproducible,fast and cheap method that does not interfere with the biofilm andthus allows further investigations with the monitored biofilm.

5. Conclusion

We found a reproducible biofilm formation of staphylococcalstandard strains (S. aureus (ATCC 29213) and S. epidermidis

RP62A (ATCC 35984)) on various bone grafts and bone graft substi-tutes under standardized in vitro conditions. The quantitative anal-ysis by a standard method (quantitative culture after washing andsonication of the material) [42] demonstrates the importance ofmonitoring factors like the specific surface or porosity of the testmaterials. Isothermal microcalorimetry proved to be a suitable toolfor an accurate, ‘‘non-invasive” and real-time microbiological assayin biofilm and material science research.

Acknowledgements

This study was supported by a research grant (E09_0001) fromthe RMS Foundation, Bettlach, Switzerland and a grant from theSwiss Orthopedic Society (SGO/SSO). Additional funding was ob-tained for the microcalorimetric analysis from the Swiss NationalFoundation (#3200B0-112547) and the Gebert Rüf Stiftung. Staph-ylococcus epidermidis RP62A (ATCC 35984) was kindly provided byProf. P. Vaudaux from the Department of Infectious Disease, Uni-versity Hospital, Geneva, Switzerland. We want to thank W. Hir-singer for his help with the SEM studies, S. Grünefelder and P.Brotschi for their help producing the samples, and Dr. A. Steinhu-ber and B. Schneider for laboratory assistance. We also thank thefollowing companies/organizations for providing the test materi-als: Biomet (Biomet Bone Cement R™), Novomedics (Tutoplast™)and RMS Foundation (b-TCP, PE).

Appendix A. Figures with essential colour discrimination

Certain figures in this article, particularly Figure 4, are difficultto interpret in black and white. The full colour images can be foundin the on-line version, at doi: 10.1016/j.actbio.2010.03.011.

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