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Room-temperature attachment of PLGA microspheres to titanium surfaces for implant-based drug release
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Room-temperature attachment of PLGA microspheres to titanium surfaces for implant-based drug release Dongqin Xiao a,b , Qing Liu a , Dongwei Wang a , Tao Xie a , Tailin Guo a , Ke Duan a,b,∗ , Jie Weng a a Key Laboratory of Advanced Technologies of Materials (Ministry of Education), School of Materials Science and Engineering, Southwest Jiaotong University, Chengdu 610031, Sichuan, China b Jiangsu Provincial Key Laboratory for Interventional Medical Devices, Huaiyin Institute of Technology, Huaiyin 223001, Jiangsu, China
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Article history: Received 23 July 2013 Received in revised form 24 April 2014 Accepted 29 April 2014 Available online xxx Keywords: Titanium Drug release PLGA Microsphere Implant surface
a b s t r a c t Drug release from implant surfaces is an effective approach to impart biological activities, (e.g., antimicrobial and osteogenic properties) to bone implants. Coatings of polylactide-based polymer are a candidate for this purpose, but a continuous (fully covering) coating may be non-optimal for implant-bone fixation. This study reports a simple room-temperature method for attaching poly (lactide-co-glycolide) (PLGA) microspheres to titanium (Ti) surfaces. Microspheres were prepared with polyvinyl alcohol (PVA) or polyvinylpyrrolidone (PVP) as the emulsifier. Microspheres were attached to Ti discs by pipetting as a suspension onto the surfaces followed by vacuum drying. After immersion in shaking water bath for 14 d, a substantial proportion of the microspheres remained attached to the discs. In contrast, if the vacuum-drying procedure was omitted, only a small fraction of the microspheres remained attached to the discs after immersion for only 5 min. Microspheres containing triclosan (a broad-spectrum antibiotic) were attached by porous-surfaced Ti discs. In vitro experiments showed that the microsphere-carrying discs were able to kill Staphylococcus aureus and Escherichia Coli, and support the adhesion and growth of primary rat osteoblasts. This simple method may offer a flexible technique for bone implant-based drug release. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Metals, such as titanium (Ti) alloys, are extensively used as bone implants thanks to their biocompatibilities. Nevertheless, current bone implants face two concerns: postoperative infection and insufficient bone formation. During implantation, bacteria may adhere to the implant surface and subsequently form a biofilm impeding drug penetration [1]. This mode of infection is difficult to eradicate by systemic antibiotic treatments and is currently the second most common cause of implant failure [2]. Additionally, in compromised circumstances (e.g., bone stock deficiency, revision surgery) new bone formation may be insufficient for achieving a reliable bone-implant fixation [3]. Under these challenging conditions, enhancement of bone formation is required. Therefore, it is
∗ Corresponding author at: School of Materials Science and Engineering, Key Lab of Advanced Technologies of Materials (MOE), Chengdu 610031, Sichuan, China. Tel.: +86 28 87601371; fax: +86 28 87601371. E-mail address: [email protected] (K. Duan).
desirable to impart biological functions to bone implants, such as antibacterial and osteogenic activities. Drug release at the bone-implant interface is an effective approach to this goal because it can attain a high local drug concentration without inducing systemic adverse effects [4]. A number of surface techniques have been developed for implant-based drug delivery [5]. Of these, lactide-based polymer coatings [e.g., polylactide (PLA) and lactide/glycolide copolymers (PLGA)] represent a promising candidate because of their advantages. A variety of drugs [6,7] can be incorporated in PLA and PLGA coatings by physical encapsulation, without strict requirements on the chemical characteristics of the drugs, such as functional groups, hydrophilicity/hydrophobicity, and molecular weight. The coatings can also serve as diffusion barriers to control the kinetics of drug release. Moreover, these polymers have been used as resorbable bone implants with positive results [8]. Studies have prepared continuous drug-loaded PLA and PLGA coatings on bone fixation wires and demonstrated their bactericidal and osteogenic activities [9–11]. However, PLA and PLGA generally lack bioactivity and cannot form a direct contact with the host bone [12,13], which is desirable for a long-term stable implant fixation. Therefore, a continuous (fully
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covering) polymer coating may be non-ideal for an implant requiring stable fixation with the host bone (e.g., artificial joints). Recent studies reported a new design to use the advantages of polymer coatings while potentially reducing the disadvantage of a continuous coating [14,15]. In this design, discrete drug-loaded PLGA microspheres are attached to Ti surfaces by a heat treatment (50 ◦ C) or supercritical carbon dioxide (CO2 ) treatment. Because microspheres do not form a continuous coating, the remaining (uncoated) Ti surface is available for direct Ti-bone contact. However, heat treatments may be non-optimal for thermally sensitive drugs (e.g., proteins) and the supercritical CO2 technique seems relatively complex and requires special equipment. Here, we report a simple method to attach PLGA microspheres to Ti surfaces by vacuum drying at room temperature. We will also demonstrate an application (in vitro bactericidal activity) of microspheres attached to Ti. 2. Materials and methods 2.1. Preparation of porous-coated Ti discs Commercially pure Ti beads ( 600 m; Honghua Nonferrous Metals, Baoji, China) were packed on Ti discs ( 10 mm × 2 mm; Zhirui Metals, Baoji, China). The bead-disc complexes were transferred into a graphite mold and sintered in vacuum (1300 ◦ C, 2 h; VMK-1800, Linn High Therm GmbH, Eschenfelden, Germany) to “weld” the beads as a porous coating on the disc [16]. 2.2. Preparation of PLGA microspheres Drug-free PLGA microspheres were prepared with polyvinyl alcohol (PVA; degree of polymerization: 1700, hydrolysis: 99%) or polyvinylpyrrolidone (PVP; Mw: 30,000, both Kelong Chemical, Chengdu, China) as the emulsifier. They are abbreviated as PLGA–PVA and PLGA–PVP, respectively. Briefly, 400 mg of PLGA (Mw: 20,000, lactide: glycolide = 75:25; Daigang Biotech, Jinan, China) was dissolved in 16 ml of dichloromethane (DCM). The solution was added dropwise into 100 ml of water containing 1 g of emulsifier (PVA or PVP). The mixture was stirred on an ice water bath (800 rpm, 10 min) to form an emulsion. After removal from the bath, the emulsion was stirred (300 rpm) for 4 h to evaporate off DCM. The formed microspheres were collected by centrifugation (4000 rpm, 5 min), rinsed with copious water (1 min × 5), and lyophilized (Advantage, SP Scientific, Gardiner, NY, USA). Microspheres containing triclosan, a broad-spectrum antibiotic (purity >99%; Xiya Reagent, Chengdu, China), were prepared by similar procedures with PVA as the emulsifier, except that 40 mg of triclosan was co-dissolved with PLGA in DCM. These microspheres are abbreviated as triclosan-PLGA–PVA. 2.3. Determination of residual emulsifier on microsphere surface Residual emulsifier on microsphere surfaces was determined by spectrophotometry. For residual PVA on PLGA-PVA microspheres [17,18], 30 mg of lyophilized microspheres were digested with 0.8 ml of 0.5 M NaOH (80 ◦ C, 20 min). The resulting liquid was neutralized with 0.36 ml of 1 M hydrochloric acid (HCl) and adjusted to 2 ml with distilled water. Then, 1.2 ml of 0.65 M boric acid, 0.2 ml of an iodine/potassium iodide solution (I2 /KI, 0.05 M/0.15 M), and 0.6 ml of distilled water were added to the liquid, forming a colored compound. After resting for 15 min, the absorbance at 650 nm was measured and converted into PVA concentration against an experimentally established standard curve (Supplementary Fig. 1a). For residual PVP on PLGA–PVP [19], 100 mg of microspheres were dissolved in 0.2 ml of DCM and then mixed with 5 ml of 0.4 M citric acid to extract PVP. The suspension was maintained at 50 ◦ C for 20 min
to evaporate off DCM and then the volume was readjusted to 5 ml. Subsequently, 1 ml of 0.006 N I2 /KI solution was added to the liquid to form a colored compound. The absorbance was measured at 500 nm and calculated into PVP concentration against a standard curve (Supplementary Fig. 1b). It is noted that, NaOH digestion was replaced by an extraction procedure in sample pretreatment for PVP quantitation, because concentrated NaOH was found to disrupt the chromogenic reaction between PVP and I2 /KI. Because of potential incomplete extraction, the extraction procedure probably led to certain underestimation of the PVP concentration in the microspheres. Nevertheless, for qualitative (or semi-quantitative) purposes, this procedure was deemed sufficient. 2.4. Attachment of drug-free microspheres to flat Ti Ti discs ( 10 mm × 2 mm) were abraded to a 1200 grit surface finish, cleaned in a mixture of nitric acid (HNO3 , 5 M) and hydrofluoric acid (HF, 0.4 M) for 30 s, and sonicated in water. Drug-free microspheres (PLGA–PVA or PLGA–PVP) were attached to the discs by two separate methods (without drying or by vacuum drying, see below) and then analyzed for the stability of attachment. Briefly, 10 mg of microspheres were dispersed in 1 ml of ultrapure water. Then, 0.1 ml of the suspension was pipetted on each disc. After standing for 5 min, several discs (i.e., attached without drying) were immediately transferred in centrifugation tubes containing 2 ml of ultrapure water and placed in a shaking water bath (37 ◦ C, 80 rpm). Other discs (i.e., attached by vacuum drying) were placed in an evacuated desiccator (25 ◦ C) for 2 h before transfer into tubes and immersion in the water bath. After immersion in the bath for predetermined periods (5 min, 2 d, 7 d, 14 d), each disc was removed and transferred in a new tube containing 2 ml of ultrapure water. The fluid in the original tube was kept for analysis. After the immersion test, all fluid samples and Ti discs were treated with 2 ml of 0.05 M sodium hydroxide (NaOH, 37 ◦ C, 48 h) to digest the contained (or attached) microspheres [20]. Digestion products were analyzed for total organic carbon contents (TOC; Vario TOC cube, Elementar GmbH, Hanau, Germany). The proportion of microspheres remaining on the disc at each time point was calculated. 2.5. Attachment of triclosan-loaded microspheres to porous-coated Ti 2.5.1. Attachment Triclosan-PLGA–PVA microspheres were attached to porouscoated Ti discs only by vacuum drying. Briefly, 15 mg of microspheres were dispersed in 1 ml of water; 0.1 ml of the suspension was pipetted on a porous-coated disc; and the disc was placed in an evacuated desiccator for 2 h. 2.5.2. In vitro triclosan release Three porous-coated Ti discs carrying triclosan-PLGA–PVA microspheres (attached as described above) were each immersed in 3 ml of phosphate buffered saline (PBS, pH 7.25) containing 0.5% (w/v) sodium dodecyl sulfate [21] (SDS, Kelong Chemical) and placed in a shaking water bath (37 ◦ C, 80 rpm) for 7 d. At predetermined intervals, 2 ml of liquid was collected for spectrophotometric analysis (281 nm, Shimadzu UV-250), and the fluid was replenished. 2.5.3. In vitro bactericidal activities The bactericidal activities of porous-coated discs carrying triclosan-PLGA–PVA microspheres were tested by co-culture with Staphylococcus aureus (ATCC 29213) and Escherichia coli (ATCC 25922; both from Guangdong Provincial Microorganism Preservation Center, Guangzhou, China). Blank porous-coated discs and
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Fig. 1. Scanning electron micrographs of (a) PLGA–PVA, (b) PLGA–PVP, and (c) triclosan-PLGA–PVA microspheres.
those carrying PLGA–PVA microspheres served as controls. Sterilized agar was poured into petri dishes and inoculated with 100 l of bacteria suspension (1.2 × 109 CFU/ml). After solidification of agar, Ti samples were gently placed on the agar. After incubation at 37 ◦ C for 48 h, the dishes were examined for the development of inhibition zones. 2.5.4. In vitro cytocompatibility The cytocompatibility of porous-coated discs carrying triclosanPLGA–PVA microspheres was evaluated using osteoblasts isolated from the calvaria of 10 day-old Sprague–Dawley rats (Experimental Animal Center, Sichuan University, Chengdu, China) [22]. After passaging for three passages, the cells were seeded on the discs at 2 × 104 cells/sample and incubated (5% CO2 , 37 ◦ C). After incubation for 1 d, 3 d, and 5 d, the cell viability was determined by Alamar Blue assay. Briefly, the medium was replaced with 800 l of Alamar Blue solution (Biosource, Camarillo, CA, USA); the culture plate was incubated at 37 ◦ C for 3 h; and the absorbance at 570 nm was measured (UQuant MQX200, Bio-Tek, Winooski, VT, USA). Porous-coated discs carrying no microspheres and those carrying PLGA–PVA microspheres served as controls. Cell-free wells served as the blank. Data were analyzed by analysis of variance (ANOVA, SPSS 11.0, SPSS, Chicago, IL, USA), and a p-value of <0.05 was considered statistically significant. 2.6. Characterizations Surface morphology was studied by scanning electron microscopy (SEM, FEI Quanta 200). Particle size distribution of microspheres was measured by laser-diffraction particle size analysis (Horiba LA-920).
immersion for 5 min (Fig. 2), only 11% of the microspheres remained on the discs. Given the poor stability of attachment without drying, no effort was made to evaluate the attachment stability after immersion for >5 min. The remarkable difference showed the essential role of vacuum drying on the stability of attachment. To probe the possible mechanisms of microsphere attachment, microspheres were synthesized using another polymer, PVP, as the emulsifier. The reasoning is as follows. PVA can be dissolved in water only at >70 ◦ C [23]. At room temperature, it is only swellable (not dissolvable) by water (after dissolution in hot water, however, PVA does not precipitate even after cooling to room temperature). Moreover, emulsifiers such as PVA can remain on the surfaces of PLGA microspheres despite repeated washing [24,25] (also see Section 3.1.2). Considering these, we hypothesized that after vacuum drying, residual PVA chains on the microsphere surfaces may have formed an insoluble thin layer “gluing” the microspheres to the Ti disc. To test this hypothesis, microspheres were prepared with PVP as the emulsifier and attached to Ti discs by vacuum drying. PVP is another frequently used emulsifier in PLGA microsphere preparation. As opposed to PVA, PVP is easily dissolved in water at room temperature. Thus, if our hypothesis is correct, PLGA–PVP microspheres should be easily detached during immersion. The PLGA–PVP microspheres (Fig. 1b) were also spherical, smooth, and ranged 3–20 m in diameter (mean diameter: 14 m). After attachment by vacuum drying and immersion in shaking water bath, PLGA–PVP microspheres stayed on Ti disc at a similar stability to PLGA–PVA microspheres. At each time point examined (Fig. 2) the proportion of PLGA–PVP microspheres remaining attached to Ti discs was similar to that recorded from PLGA–PVA
3. Results and discussion 3.1. Microsphere attachment on flat Ti 3.1.1. Attachment stability The PLGA–PVA microspheres (Fig. 1a) were spherical, smooth, and 5–25 m in diameter (mean diameter: 18 m). Serendipitously, we observed that after dispense as a suspension on Ti followed by vacuum drying, these microspheres could remain on the surface even after immersion in water for a few weeks. A quantitative immersion experiment was thus performed to evaluate the stability of this attachment. TOC assays (Fig. 2) found that, when PLGA–PVA microspheres were attached to flat Ti discs by vacuum drying, after subsequent immersion in shaking water bath for 5 min, 93% of the microspheres remained attached to the discs. The proportion of microspheres remaining attached to the discs decreased gradually with time. After immersion for 14 d, 67% of the microspheres remained on the discs. In comparison, when microspheres were simply dispensed as a suspension on Ti discs without vacuum drying, after subsequent
Fig. 2. Proportion of PLGA–PVA microspheres remaining attached to Ti discs after immersion in shaking water bath for various periods (mean ± standard deviation, n = 3).
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Fig. 3. Scanning electron micrographs showing PLGA microspheres attached to flat Ti discs after immersion in shaking water bath for various periods.
microspheres attached by vacuum drying. This result indicated that emulsifier insolubility is not essential for a stable microsphere attachment. SEM studies were performed to gain more insights into the attached microspheres during immersion in the water bath. When microspheres were attached by vacuum drying, the phenomena (Fig. 3, top and middle rows) were similar regardless of the type of emulsifier used. After immersion for 5 min, microspheres partially covered the disc, and a fraction of microspheres appeared deposited on top of underlying ones instead of directly attached to the disc. After immersion for 2 d, the surface morphology was largely similar. After immersion for 14 d, adjacent microspheres partially coalesced and flattened, presumably because of the low glass transition temperature of the PLGA used in this study (∼36 ◦ C). When microspheres were attached without drying (Fig. 3 bottom row), after immersion for 5 min, fewer microspheres remained on the disc. More, the microspheres remaining attached were substantially smaller than those attached by vacuum drying, probably because only smaller microspheres could be attached (greater surface/volume ratios) when limited microsphere-substrate bonds (interactions) were formed.
3.1.2. Residual emulsifier on microsphere surface Spectrophotometric analyses found that, the PLGA–PVA microspheres contained 0.16 ± 0.01% (w/w) residual PVA, and PLGA–PVP contained 0.05 ± 0.01% (w/w) residual PVP. The residual PVA concentration was of the same order of magnitude to the lower range (i.e., 0.7%) reported for PLGA micro-particles [17] and at least one order of magnitude lower than values determined from PLGA nano-particles [18]. The differences may be attributed to different diameters and PVA concentrations used in microsphere preparation. The residual PVP concentration could not be compared with literature results because of the lack of quantitative works on this emulsifier.
3.1.3. Attachment mechanisms The exact mechanisms of microsphere attachment to Ti discs are not fully understood, but a preliminary explanation may be suggested. A fraction of emulsifier (PVA or PVP) chains remained on the microsphere surfaces despite repeated rinsing [24,25]. Shortly after dispense as a suspension on Ti, these chains became in contact with the Ti surface. As PVA and PVP are both adhesives, their side groups formed local bonds with Ti, such as coordination [26] and acid–base interaction [27]. Without drying, the chains were solvated by water, and the Ti surface was also wetted by a thin film of water. As a result, only a small number of such bonds could form; the microspheres and Ti were largely separated by water; and the microsphere attachment was relatively weak. In comparison, when water was removed by vacuum drying, the chains were forced to form more local contacts with Ti, thereby creating more bonds and increasing the attachment strength. Moreover, polar interactions between Ti and carbonyl groups in PLGA as well as mechanical interlocks may have also contributed to the attachment. At the later stage of immersion, microsphere flattening may have also facilitated the microsphere attachment (i.e., increased surface/volume ratio). The relative contribution of each attachment force, however, is unknown.
3.2. Microsphere attachment to porous-coated Ti 3.2.1. Attachment to discs Despite inadequate understanding of the attachment mechanisms, this reasonably stable attachment offered a simple method of imparting biological functions to Ti. As a demonstration, triclosan-loaded PLGA microspheres were prepared and attached to porous-coated Ti discs by vacuum drying. Triclosan is a broadspectrum antibiotic against both Gram-positive and Gram-negative bacteria. This drug has also been applied on medical devices for the prevention of postoperative infections [28,29].
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Fig. 4. Scanning electron micrographs of (a) an as-sintered porous-coated Ti disc and (b) and (c) a porous-coated Ti-disc carrying triclosan-PLGA–PVA microspheres attached by vacuum drying (arrows point to microspheres).
The porous-coated discs (Fig. 4a) had pores 150–500 m in size and an estimated porosity of 37%, similar to the surfaces of actual bone implants [16]. The triclosan-PLGA–PVA microspheres were spherical, smooth, and 2–20 m in diameter (mean diameter: 12 m). Spectrophotometric assays confirmed that the microspheres contained 5.5 wt% of triclosan. After attachment by vacuum drying (Fig. 4b and c), microspheres were deposited on Ti beads covering a fraction of the bead surface. Some microspheres were concentrated at junction areas between beads (Fig. 4c), but the pores between beads generally remained open (Fig. 4b).
3.2.2. In vitro performance The porous-coated discs carrying triclosan-PLGA–PVA microspheres showed a burst release (∼80%) (Fig. 5) of triclosan in the first 10 h followed by a slow release for up to 5 d. Because triclosan is poorly soluble in water, its release into organic-free PBS was slow and the concentration could not be accurately determined by UV spectrophotometry. Therefore, SDS (a solubilizing agent) was added in the medium to facilitate triclosan release. This practice has been commonly used in evaluating triclosan release from microspheres and coatings [21,30]. Although SDS is absent in the physiological environment, it may be argued that biological molecules (e.g., lipids) in vivo may act as solubilizing agents for triclosan.
Fig. 5. In vitro release of triclosan from porous-coated Ti discs carrying triclosanPLGA–PVA microspheres attached by vacuum drying (mean ± standard deviation, n = 3).
Earlier studies have identified the first 6 h after device implantation to be critical for preventing implant-related infections [31,32]. During this “decisive period”, the implant is particularly susceptible to bacterial colonization [31]. This period is also the prime
Fig. 6. In vitro bactericidal activities of porous-coated Ti discs carrying no microspheres or carrying PLGA microspheres against S. aureus and E. coli.
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Fig. 8. (a) Light and (b) scanning electron micrographs showing a square pattern of PLGA–PVA microspheres formed on a Ti disc using a punched Parafilm as the mask.
Fig. 7. Viability of primary rat osteoblasts seeded on blank porous-coated Ti discs, discs carrying PLGA–PVA microspheres, and those carrying triclosan-PLGA–PVA microspheres after culture for various periods (Alamar Blue assay; mean ± standard deviation, n = 3).
opportunity to eradicate pathogens at the implant site [32]. Accordingly, Vasilev et al. [33] suggested that an ideal antibiotic-release coating should give a fast initial release for 6 h followed by a sustained slower release. The triclosan release from the porous-coated Ti (Fig. 5) appeared to generally meet this requirement, and thus is potentially suitable for preventing postoperative infection. However, modification of release rate may be required for meeting various complicated clinical situations (e.g., revision of artificial joints with known/suspected history of infection). After co-culture with S. aureus and E. coli for 48 h (Fig. 6), the discs carrying triclosan-PLGA–PVA microspheres produced clear zones of inhibition against S. aureus (diameter: 43.4 ± 1.5 mm) and E. coli (diameter: 30.5 ± 2.8 mm, both n = 3). In comparison, discs carrying no microspheres or (drug-free) PLGA–PVA microspheres formed no inhibition zones. S. aureus is a Gram-positive bacterium and is also a major pathogen responsible for implant infection [31]. E. coli is a Gram-negative bacterium and has been used as a model pathogen in evaluating the antimicrobial activities of bone implant surfaces [34]. These results suggested that, the microsphere-carrying discs were effective at killing adjacent bacteria. The cytocompatibility of the microsphere-carrying discs were examined by culture with primary rat osteoblasts. Alamar Blue assays (Fig. 7) showed that, all groups supported cell proliferation up to day 5 and no statistically significant difference was found at any time point examined (all p > 0.05). These results suggested that the microsphere-carrying discs were safe and biocompatible with osteoblasts.
of various drugs with relative few requirements on their physicochemical properties (e.g., water- or oil-soluble, functional groups, negatively or positively charged). Moreover, it does not form a continuous coating and, thus, allows direct bone-implant contact. However, the method may involve two potential concerns. First, as a result of microsphere agglomeration during drying, the microsphere distribution on the substrate tends to be non-uniform. We conducted a preliminary experiment to control microsphere distribution on a planar substrate by a simple patterning method [35]. Briefly, a Parafilm (Bemis, Neenah, WI, USA) was punched with a blunt-ended needle to create a square pattern of holes ( 400 m). After firmly pressing the Parafilm onto a Ti disc (1200-grit surface finish), a suspension of PLGA–PVA microspheres (10 mg/ml) was injected to each hole using a 1-ml syringe with a 25 ga needle; the sample was vacuum-dried for 1.5 h; and the Parafilm was carefully removed. Light and electron microscopy (Fig. 8) showed that, microspheres formed a regular array of “islands” on the substrate. Although obviously immature at present, this result suggests the possibility to control microsphere distribution on substrates. With the help of more advanced techniques, such as microcontact printing [36] and ink-jet microprinting [37], it may be possible to precisely control microsphere distribution on substrates. Another concern of this method is the occupation of a fraction of the implant surface (or pore space) available for bone contact (or ingrowth). This may be alleviated by reducing the amount of microspheres need to be attached (i.e., loading of potent drugs) and distributing the microspheres in deeper pores. Recent experiments suggested that this attachment method is applicable to other substrates such as hydroxyapatite. It would be interesting to investigate whether the method is suitable for microspheres of other materials. 4. Conclusion PLGA microspheres synthesized with PVA or PVP as the emulsifier were attached to Ti surfaces by a simple vacuum drying method. This attachment is not reliant on the emulsifier insolubility. A substantial proportion of the microspheres remained attached to Ti discs after immersion in water bath for 2 weeks. Triclosanloaded PLGA microspheres were attached to porous-coated Ti discs. The resulting microsphere-carrying discs were able to kill bacteria in vitro and allow rat osteoblast adhesion and growth. This simple room-temperature method offers a flexible technique for drug release from bone implant surfaces. Future studies are needed to determine whether this method can be applied to other substrates and microspheres. Acknowledgments
3.3. Microsphere attachment as a general drug-release surface design This simple room-temperature method provides a flexible technique for implant-based drug release, allowing controlled release
This study was supported by National Basic Research Program of China (973 Program, 2012CB933600), National Natural Science Foundation of China (51172188, 81071456), National Engineering Research Centerfor Tissue Restoration and Reconstruction
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Room-temperature attachment of PLGA microspheres to titanium surfaces for implant-based drug release Dongqin Xiao a,b , Qing Liu a , Dongwei Wang a , Tao Xie a , Tailin Guo a , Ke Duan a,b,∗ , Jie Weng a a Key Laboratory of Advanced Technologies of Materials (Ministry of Education), School of Materials Science and Engineering, Southwest Jiaotong University, Chengdu 610031, Sichuan, China b Jiangsu Provincial Key Laboratory for Interventional Medical Devices, Huaiyin Institute of Technology, Huaiyin 223001, Jiangsu, China
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Article history: Received 23 July 2013 Received in revised form 24 April 2014 Accepted 29 April 2014 Available online xxx Keywords: Titanium Drug release PLGA Microsphere Implant surface
a b s t r a c t Drug release from implant surfaces is an effective approach to impart biological activities, (e.g., antimicrobial and osteogenic properties) to bone implants. Coatings of polylactide-based polymer are a candidate for this purpose, but a continuous (fully covering) coating may be non-optimal for implant-bone fixation. This study reports a simple room-temperature method for attaching poly (lactide-co-glycolide) (PLGA) microspheres to titanium (Ti) surfaces. Microspheres were prepared with polyvinyl alcohol (PVA) or polyvinylpyrrolidone (PVP) as the emulsifier. Microspheres were attached to Ti discs by pipetting as a suspension onto the surfaces followed by vacuum drying. After immersion in shaking water bath for 14 d, a substantial proportion of the microspheres remained attached to the discs. In contrast, if the vacuum-drying procedure was omitted, only a small fraction of the microspheres remained attached to the discs after immersion for only 5 min. Microspheres containing triclosan (a broad-spectrum antibiotic) were attached by porous-surfaced Ti discs. In vitro experiments showed that the microsphere-carrying discs were able to kill Staphylococcus aureus and Escherichia Coli, and support the adhesion and growth of primary rat osteoblasts. This simple method may offer a flexible technique for bone implant-based drug release. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Metals, such as titanium (Ti) alloys, are extensively used as bone implants thanks to their biocompatibilities. Nevertheless, current bone implants face two concerns: postoperative infection and insufficient bone formation. During implantation, bacteria may adhere to the implant surface and subsequently form a biofilm impeding drug penetration [1]. This mode of infection is difficult to eradicate by systemic antibiotic treatments and is currently the second most common cause of implant failure [2]. Additionally, in compromised circumstances (e.g., bone stock deficiency, revision surgery) new bone formation may be insufficient for achieving a reliable bone-implant fixation [3]. Under these challenging conditions, enhancement of bone formation is required. Therefore, it is
∗ Corresponding author at: School of Materials Science and Engineering, Key Lab of Advanced Technologies of Materials (MOE), Chengdu 610031, Sichuan, China. Tel.: +86 28 87601371; fax: +86 28 87601371. E-mail address: [email protected] (K. Duan).
desirable to impart biological functions to bone implants, such as antibacterial and osteogenic activities. Drug release at the bone-implant interface is an effective approach to this goal because it can attain a high local drug concentration without inducing systemic adverse effects [4]. A number of surface techniques have been developed for implant-based drug delivery [5]. Of these, lactide-based polymer coatings [e.g., polylactide (PLA) and lactide/glycolide copolymers (PLGA)] represent a promising candidate because of their advantages. A variety of drugs [6,7] can be incorporated in PLA and PLGA coatings by physical encapsulation, without strict requirements on the chemical characteristics of the drugs, such as functional groups, hydrophilicity/hydrophobicity, and molecular weight. The coatings can also serve as diffusion barriers to control the kinetics of drug release. Moreover, these polymers have been used as resorbable bone implants with positive results [8]. Studies have prepared continuous drug-loaded PLA and PLGA coatings on bone fixation wires and demonstrated their bactericidal and osteogenic activities [9–11]. However, PLA and PLGA generally lack bioactivity and cannot form a direct contact with the host bone [12,13], which is desirable for a long-term stable implant fixation. Therefore, a continuous (fully
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covering) polymer coating may be non-ideal for an implant requiring stable fixation with the host bone (e.g., artificial joints). Recent studies reported a new design to use the advantages of polymer coatings while potentially reducing the disadvantage of a continuous coating [14,15]. In this design, discrete drug-loaded PLGA microspheres are attached to Ti surfaces by a heat treatment (50 ◦ C) or supercritical carbon dioxide (CO2 ) treatment. Because microspheres do not form a continuous coating, the remaining (uncoated) Ti surface is available for direct Ti-bone contact. However, heat treatments may be non-optimal for thermally sensitive drugs (e.g., proteins) and the supercritical CO2 technique seems relatively complex and requires special equipment. Here, we report a simple method to attach PLGA microspheres to Ti surfaces by vacuum drying at room temperature. We will also demonstrate an application (in vitro bactericidal activity) of microspheres attached to Ti. 2. Materials and methods 2.1. Preparation of porous-coated Ti discs Commercially pure Ti beads ( 600 m; Honghua Nonferrous Metals, Baoji, China) were packed on Ti discs ( 10 mm × 2 mm; Zhirui Metals, Baoji, China). The bead-disc complexes were transferred into a graphite mold and sintered in vacuum (1300 ◦ C, 2 h; VMK-1800, Linn High Therm GmbH, Eschenfelden, Germany) to “weld” the beads as a porous coating on the disc [16]. 2.2. Preparation of PLGA microspheres Drug-free PLGA microspheres were prepared with polyvinyl alcohol (PVA; degree of polymerization: 1700, hydrolysis: 99%) or polyvinylpyrrolidone (PVP; Mw: 30,000, both Kelong Chemical, Chengdu, China) as the emulsifier. They are abbreviated as PLGA–PVA and PLGA–PVP, respectively. Briefly, 400 mg of PLGA (Mw: 20,000, lactide: glycolide = 75:25; Daigang Biotech, Jinan, China) was dissolved in 16 ml of dichloromethane (DCM). The solution was added dropwise into 100 ml of water containing 1 g of emulsifier (PVA or PVP). The mixture was stirred on an ice water bath (800 rpm, 10 min) to form an emulsion. After removal from the bath, the emulsion was stirred (300 rpm) for 4 h to evaporate off DCM. The formed microspheres were collected by centrifugation (4000 rpm, 5 min), rinsed with copious water (1 min × 5), and lyophilized (Advantage, SP Scientific, Gardiner, NY, USA). Microspheres containing triclosan, a broad-spectrum antibiotic (purity >99%; Xiya Reagent, Chengdu, China), were prepared by similar procedures with PVA as the emulsifier, except that 40 mg of triclosan was co-dissolved with PLGA in DCM. These microspheres are abbreviated as triclosan-PLGA–PVA. 2.3. Determination of residual emulsifier on microsphere surface Residual emulsifier on microsphere surfaces was determined by spectrophotometry. For residual PVA on PLGA-PVA microspheres [17,18], 30 mg of lyophilized microspheres were digested with 0.8 ml of 0.5 M NaOH (80 ◦ C, 20 min). The resulting liquid was neutralized with 0.36 ml of 1 M hydrochloric acid (HCl) and adjusted to 2 ml with distilled water. Then, 1.2 ml of 0.65 M boric acid, 0.2 ml of an iodine/potassium iodide solution (I2 /KI, 0.05 M/0.15 M), and 0.6 ml of distilled water were added to the liquid, forming a colored compound. After resting for 15 min, the absorbance at 650 nm was measured and converted into PVA concentration against an experimentally established standard curve (Supplementary Fig. 1a). For residual PVP on PLGA–PVP [19], 100 mg of microspheres were dissolved in 0.2 ml of DCM and then mixed with 5 ml of 0.4 M citric acid to extract PVP. The suspension was maintained at 50 ◦ C for 20 min
to evaporate off DCM and then the volume was readjusted to 5 ml. Subsequently, 1 ml of 0.006 N I2 /KI solution was added to the liquid to form a colored compound. The absorbance was measured at 500 nm and calculated into PVP concentration against a standard curve (Supplementary Fig. 1b). It is noted that, NaOH digestion was replaced by an extraction procedure in sample pretreatment for PVP quantitation, because concentrated NaOH was found to disrupt the chromogenic reaction between PVP and I2 /KI. Because of potential incomplete extraction, the extraction procedure probably led to certain underestimation of the PVP concentration in the microspheres. Nevertheless, for qualitative (or semi-quantitative) purposes, this procedure was deemed sufficient. 2.4. Attachment of drug-free microspheres to flat Ti Ti discs ( 10 mm × 2 mm) were abraded to a 1200 grit surface finish, cleaned in a mixture of nitric acid (HNO3 , 5 M) and hydrofluoric acid (HF, 0.4 M) for 30 s, and sonicated in water. Drug-free microspheres (PLGA–PVA or PLGA–PVP) were attached to the discs by two separate methods (without drying or by vacuum drying, see below) and then analyzed for the stability of attachment. Briefly, 10 mg of microspheres were dispersed in 1 ml of ultrapure water. Then, 0.1 ml of the suspension was pipetted on each disc. After standing for 5 min, several discs (i.e., attached without drying) were immediately transferred in centrifugation tubes containing 2 ml of ultrapure water and placed in a shaking water bath (37 ◦ C, 80 rpm). Other discs (i.e., attached by vacuum drying) were placed in an evacuated desiccator (25 ◦ C) for 2 h before transfer into tubes and immersion in the water bath. After immersion in the bath for predetermined periods (5 min, 2 d, 7 d, 14 d), each disc was removed and transferred in a new tube containing 2 ml of ultrapure water. The fluid in the original tube was kept for analysis. After the immersion test, all fluid samples and Ti discs were treated with 2 ml of 0.05 M sodium hydroxide (NaOH, 37 ◦ C, 48 h) to digest the contained (or attached) microspheres [20]. Digestion products were analyzed for total organic carbon contents (TOC; Vario TOC cube, Elementar GmbH, Hanau, Germany). The proportion of microspheres remaining on the disc at each time point was calculated. 2.5. Attachment of triclosan-loaded microspheres to porous-coated Ti 2.5.1. Attachment Triclosan-PLGA–PVA microspheres were attached to porouscoated Ti discs only by vacuum drying. Briefly, 15 mg of microspheres were dispersed in 1 ml of water; 0.1 ml of the suspension was pipetted on a porous-coated disc; and the disc was placed in an evacuated desiccator for 2 h. 2.5.2. In vitro triclosan release Three porous-coated Ti discs carrying triclosan-PLGA–PVA microspheres (attached as described above) were each immersed in 3 ml of phosphate buffered saline (PBS, pH 7.25) containing 0.5% (w/v) sodium dodecyl sulfate [21] (SDS, Kelong Chemical) and placed in a shaking water bath (37 ◦ C, 80 rpm) for 7 d. At predetermined intervals, 2 ml of liquid was collected for spectrophotometric analysis (281 nm, Shimadzu UV-250), and the fluid was replenished. 2.5.3. In vitro bactericidal activities The bactericidal activities of porous-coated discs carrying triclosan-PLGA–PVA microspheres were tested by co-culture with Staphylococcus aureus (ATCC 29213) and Escherichia coli (ATCC 25922; both from Guangdong Provincial Microorganism Preservation Center, Guangzhou, China). Blank porous-coated discs and
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Fig. 1. Scanning electron micrographs of (a) PLGA–PVA, (b) PLGA–PVP, and (c) triclosan-PLGA–PVA microspheres.
those carrying PLGA–PVA microspheres served as controls. Sterilized agar was poured into petri dishes and inoculated with 100 l of bacteria suspension (1.2 × 109 CFU/ml). After solidification of agar, Ti samples were gently placed on the agar. After incubation at 37 ◦ C for 48 h, the dishes were examined for the development of inhibition zones. 2.5.4. In vitro cytocompatibility The cytocompatibility of porous-coated discs carrying triclosanPLGA–PVA microspheres was evaluated using osteoblasts isolated from the calvaria of 10 day-old Sprague–Dawley rats (Experimental Animal Center, Sichuan University, Chengdu, China) [22]. After passaging for three passages, the cells were seeded on the discs at 2 × 104 cells/sample and incubated (5% CO2 , 37 ◦ C). After incubation for 1 d, 3 d, and 5 d, the cell viability was determined by Alamar Blue assay. Briefly, the medium was replaced with 800 l of Alamar Blue solution (Biosource, Camarillo, CA, USA); the culture plate was incubated at 37 ◦ C for 3 h; and the absorbance at 570 nm was measured (UQuant MQX200, Bio-Tek, Winooski, VT, USA). Porous-coated discs carrying no microspheres and those carrying PLGA–PVA microspheres served as controls. Cell-free wells served as the blank. Data were analyzed by analysis of variance (ANOVA, SPSS 11.0, SPSS, Chicago, IL, USA), and a p-value of <0.05 was considered statistically significant. 2.6. Characterizations Surface morphology was studied by scanning electron microscopy (SEM, FEI Quanta 200). Particle size distribution of microspheres was measured by laser-diffraction particle size analysis (Horiba LA-920).
immersion for 5 min (Fig. 2), only 11% of the microspheres remained on the discs. Given the poor stability of attachment without drying, no effort was made to evaluate the attachment stability after immersion for >5 min. The remarkable difference showed the essential role of vacuum drying on the stability of attachment. To probe the possible mechanisms of microsphere attachment, microspheres were synthesized using another polymer, PVP, as the emulsifier. The reasoning is as follows. PVA can be dissolved in water only at >70 ◦ C [23]. At room temperature, it is only swellable (not dissolvable) by water (after dissolution in hot water, however, PVA does not precipitate even after cooling to room temperature). Moreover, emulsifiers such as PVA can remain on the surfaces of PLGA microspheres despite repeated washing [24,25] (also see Section 3.1.2). Considering these, we hypothesized that after vacuum drying, residual PVA chains on the microsphere surfaces may have formed an insoluble thin layer “gluing” the microspheres to the Ti disc. To test this hypothesis, microspheres were prepared with PVP as the emulsifier and attached to Ti discs by vacuum drying. PVP is another frequently used emulsifier in PLGA microsphere preparation. As opposed to PVA, PVP is easily dissolved in water at room temperature. Thus, if our hypothesis is correct, PLGA–PVP microspheres should be easily detached during immersion. The PLGA–PVP microspheres (Fig. 1b) were also spherical, smooth, and ranged 3–20 m in diameter (mean diameter: 14 m). After attachment by vacuum drying and immersion in shaking water bath, PLGA–PVP microspheres stayed on Ti disc at a similar stability to PLGA–PVA microspheres. At each time point examined (Fig. 2) the proportion of PLGA–PVP microspheres remaining attached to Ti discs was similar to that recorded from PLGA–PVA
3. Results and discussion 3.1. Microsphere attachment on flat Ti 3.1.1. Attachment stability The PLGA–PVA microspheres (Fig. 1a) were spherical, smooth, and 5–25 m in diameter (mean diameter: 18 m). Serendipitously, we observed that after dispense as a suspension on Ti followed by vacuum drying, these microspheres could remain on the surface even after immersion in water for a few weeks. A quantitative immersion experiment was thus performed to evaluate the stability of this attachment. TOC assays (Fig. 2) found that, when PLGA–PVA microspheres were attached to flat Ti discs by vacuum drying, after subsequent immersion in shaking water bath for 5 min, 93% of the microspheres remained attached to the discs. The proportion of microspheres remaining attached to the discs decreased gradually with time. After immersion for 14 d, 67% of the microspheres remained on the discs. In comparison, when microspheres were simply dispensed as a suspension on Ti discs without vacuum drying, after subsequent
Fig. 2. Proportion of PLGA–PVA microspheres remaining attached to Ti discs after immersion in shaking water bath for various periods (mean ± standard deviation, n = 3).
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Fig. 3. Scanning electron micrographs showing PLGA microspheres attached to flat Ti discs after immersion in shaking water bath for various periods.
microspheres attached by vacuum drying. This result indicated that emulsifier insolubility is not essential for a stable microsphere attachment. SEM studies were performed to gain more insights into the attached microspheres during immersion in the water bath. When microspheres were attached by vacuum drying, the phenomena (Fig. 3, top and middle rows) were similar regardless of the type of emulsifier used. After immersion for 5 min, microspheres partially covered the disc, and a fraction of microspheres appeared deposited on top of underlying ones instead of directly attached to the disc. After immersion for 2 d, the surface morphology was largely similar. After immersion for 14 d, adjacent microspheres partially coalesced and flattened, presumably because of the low glass transition temperature of the PLGA used in this study (∼36 ◦ C). When microspheres were attached without drying (Fig. 3 bottom row), after immersion for 5 min, fewer microspheres remained on the disc. More, the microspheres remaining attached were substantially smaller than those attached by vacuum drying, probably because only smaller microspheres could be attached (greater surface/volume ratios) when limited microsphere-substrate bonds (interactions) were formed.
3.1.2. Residual emulsifier on microsphere surface Spectrophotometric analyses found that, the PLGA–PVA microspheres contained 0.16 ± 0.01% (w/w) residual PVA, and PLGA–PVP contained 0.05 ± 0.01% (w/w) residual PVP. The residual PVA concentration was of the same order of magnitude to the lower range (i.e., 0.7%) reported for PLGA micro-particles [17] and at least one order of magnitude lower than values determined from PLGA nano-particles [18]. The differences may be attributed to different diameters and PVA concentrations used in microsphere preparation. The residual PVP concentration could not be compared with literature results because of the lack of quantitative works on this emulsifier.
3.1.3. Attachment mechanisms The exact mechanisms of microsphere attachment to Ti discs are not fully understood, but a preliminary explanation may be suggested. A fraction of emulsifier (PVA or PVP) chains remained on the microsphere surfaces despite repeated rinsing [24,25]. Shortly after dispense as a suspension on Ti, these chains became in contact with the Ti surface. As PVA and PVP are both adhesives, their side groups formed local bonds with Ti, such as coordination [26] and acid–base interaction [27]. Without drying, the chains were solvated by water, and the Ti surface was also wetted by a thin film of water. As a result, only a small number of such bonds could form; the microspheres and Ti were largely separated by water; and the microsphere attachment was relatively weak. In comparison, when water was removed by vacuum drying, the chains were forced to form more local contacts with Ti, thereby creating more bonds and increasing the attachment strength. Moreover, polar interactions between Ti and carbonyl groups in PLGA as well as mechanical interlocks may have also contributed to the attachment. At the later stage of immersion, microsphere flattening may have also facilitated the microsphere attachment (i.e., increased surface/volume ratio). The relative contribution of each attachment force, however, is unknown.
3.2. Microsphere attachment to porous-coated Ti 3.2.1. Attachment to discs Despite inadequate understanding of the attachment mechanisms, this reasonably stable attachment offered a simple method of imparting biological functions to Ti. As a demonstration, triclosan-loaded PLGA microspheres were prepared and attached to porous-coated Ti discs by vacuum drying. Triclosan is a broadspectrum antibiotic against both Gram-positive and Gram-negative bacteria. This drug has also been applied on medical devices for the prevention of postoperative infections [28,29].
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Fig. 4. Scanning electron micrographs of (a) an as-sintered porous-coated Ti disc and (b) and (c) a porous-coated Ti-disc carrying triclosan-PLGA–PVA microspheres attached by vacuum drying (arrows point to microspheres).
The porous-coated discs (Fig. 4a) had pores 150–500 m in size and an estimated porosity of 37%, similar to the surfaces of actual bone implants [16]. The triclosan-PLGA–PVA microspheres were spherical, smooth, and 2–20 m in diameter (mean diameter: 12 m). Spectrophotometric assays confirmed that the microspheres contained 5.5 wt% of triclosan. After attachment by vacuum drying (Fig. 4b and c), microspheres were deposited on Ti beads covering a fraction of the bead surface. Some microspheres were concentrated at junction areas between beads (Fig. 4c), but the pores between beads generally remained open (Fig. 4b).
3.2.2. In vitro performance The porous-coated discs carrying triclosan-PLGA–PVA microspheres showed a burst release (∼80%) (Fig. 5) of triclosan in the first 10 h followed by a slow release for up to 5 d. Because triclosan is poorly soluble in water, its release into organic-free PBS was slow and the concentration could not be accurately determined by UV spectrophotometry. Therefore, SDS (a solubilizing agent) was added in the medium to facilitate triclosan release. This practice has been commonly used in evaluating triclosan release from microspheres and coatings [21,30]. Although SDS is absent in the physiological environment, it may be argued that biological molecules (e.g., lipids) in vivo may act as solubilizing agents for triclosan.
Fig. 5. In vitro release of triclosan from porous-coated Ti discs carrying triclosanPLGA–PVA microspheres attached by vacuum drying (mean ± standard deviation, n = 3).
Earlier studies have identified the first 6 h after device implantation to be critical for preventing implant-related infections [31,32]. During this “decisive period”, the implant is particularly susceptible to bacterial colonization [31]. This period is also the prime
Fig. 6. In vitro bactericidal activities of porous-coated Ti discs carrying no microspheres or carrying PLGA microspheres against S. aureus and E. coli.
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Fig. 8. (a) Light and (b) scanning electron micrographs showing a square pattern of PLGA–PVA microspheres formed on a Ti disc using a punched Parafilm as the mask.
Fig. 7. Viability of primary rat osteoblasts seeded on blank porous-coated Ti discs, discs carrying PLGA–PVA microspheres, and those carrying triclosan-PLGA–PVA microspheres after culture for various periods (Alamar Blue assay; mean ± standard deviation, n = 3).
opportunity to eradicate pathogens at the implant site [32]. Accordingly, Vasilev et al. [33] suggested that an ideal antibiotic-release coating should give a fast initial release for 6 h followed by a sustained slower release. The triclosan release from the porous-coated Ti (Fig. 5) appeared to generally meet this requirement, and thus is potentially suitable for preventing postoperative infection. However, modification of release rate may be required for meeting various complicated clinical situations (e.g., revision of artificial joints with known/suspected history of infection). After co-culture with S. aureus and E. coli for 48 h (Fig. 6), the discs carrying triclosan-PLGA–PVA microspheres produced clear zones of inhibition against S. aureus (diameter: 43.4 ± 1.5 mm) and E. coli (diameter: 30.5 ± 2.8 mm, both n = 3). In comparison, discs carrying no microspheres or (drug-free) PLGA–PVA microspheres formed no inhibition zones. S. aureus is a Gram-positive bacterium and is also a major pathogen responsible for implant infection [31]. E. coli is a Gram-negative bacterium and has been used as a model pathogen in evaluating the antimicrobial activities of bone implant surfaces [34]. These results suggested that, the microsphere-carrying discs were effective at killing adjacent bacteria. The cytocompatibility of the microsphere-carrying discs were examined by culture with primary rat osteoblasts. Alamar Blue assays (Fig. 7) showed that, all groups supported cell proliferation up to day 5 and no statistically significant difference was found at any time point examined (all p > 0.05). These results suggested that the microsphere-carrying discs were safe and biocompatible with osteoblasts.
of various drugs with relative few requirements on their physicochemical properties (e.g., water- or oil-soluble, functional groups, negatively or positively charged). Moreover, it does not form a continuous coating and, thus, allows direct bone-implant contact. However, the method may involve two potential concerns. First, as a result of microsphere agglomeration during drying, the microsphere distribution on the substrate tends to be non-uniform. We conducted a preliminary experiment to control microsphere distribution on a planar substrate by a simple patterning method [35]. Briefly, a Parafilm (Bemis, Neenah, WI, USA) was punched with a blunt-ended needle to create a square pattern of holes ( 400 m). After firmly pressing the Parafilm onto a Ti disc (1200-grit surface finish), a suspension of PLGA–PVA microspheres (10 mg/ml) was injected to each hole using a 1-ml syringe with a 25 ga needle; the sample was vacuum-dried for 1.5 h; and the Parafilm was carefully removed. Light and electron microscopy (Fig. 8) showed that, microspheres formed a regular array of “islands” on the substrate. Although obviously immature at present, this result suggests the possibility to control microsphere distribution on substrates. With the help of more advanced techniques, such as microcontact printing [36] and ink-jet microprinting [37], it may be possible to precisely control microsphere distribution on substrates. Another concern of this method is the occupation of a fraction of the implant surface (or pore space) available for bone contact (or ingrowth). This may be alleviated by reducing the amount of microspheres need to be attached (i.e., loading of potent drugs) and distributing the microspheres in deeper pores. Recent experiments suggested that this attachment method is applicable to other substrates such as hydroxyapatite. It would be interesting to investigate whether the method is suitable for microspheres of other materials. 4. Conclusion PLGA microspheres synthesized with PVA or PVP as the emulsifier were attached to Ti surfaces by a simple vacuum drying method. This attachment is not reliant on the emulsifier insolubility. A substantial proportion of the microspheres remained attached to Ti discs after immersion in water bath for 2 weeks. Triclosanloaded PLGA microspheres were attached to porous-coated Ti discs. The resulting microsphere-carrying discs were able to kill bacteria in vitro and allow rat osteoblast adhesion and growth. This simple room-temperature method offers a flexible technique for drug release from bone implant surfaces. Future studies are needed to determine whether this method can be applied to other substrates and microspheres. Acknowledgments
3.3. Microsphere attachment as a general drug-release surface design This simple room-temperature method provides a flexible technique for implant-based drug release, allowing controlled release
This study was supported by National Basic Research Program of China (973 Program, 2012CB933600), National Natural Science Foundation of China (51172188, 81071456), National Engineering Research Centerfor Tissue Restoration and Reconstruction
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Please cite this article in press as: D. Xiao, et al., Room-temperature attachment of PLGA microspheres to titanium surfaces for implantbased drug release, Appl. Surf. Sci. (2014), http://dx.doi.org/10.1016/j.apsusc.2014.04.195