[Skip to Content]
[Skip to Content Landing]
Figure.
Representative Scanning Electron Microscopy Images of Facial Implant Samples
Representative Scanning Electron Microscopy Images of Facial Implant Samples

A, Nasal dorsal silicone implant from patient 1 (original magnification ×500). B, Nasal dorsal porous polyethylene implant from patient 2 (original magnification ×1200). Information about patients 1 and 2 are given in the Table.

Table.  
Summary of the Presence of Biofilm Formationa
Summary of the Presence of Biofilm Formationa
1.
Costerton  JW, Stewart  PS, Greenberg  EP.  Bacterial biofilms: a common cause of persistent infections.  Science. 1999;284(5418):1318-1322.PubMedArticle
2.
Stoodley  P, Sauer  K, Davies  DG, Costerton  JW.  Biofilms as complex differentiated communities.  Annu Rev Microbiol. 2002;56:187-209.PubMedArticle
3.
Hall-Stoodley  L, Costerton  JW, Stoodley  P.  Bacterial biofilms: from the natural environment to infectious diseases.  Nat Rev Microbiol. 2004;2(2):95-108.PubMedArticle
4.
Ribeiro  M, Monteiro  FJ, Ferraz  MP.  Infection of orthopedic implants with emphasis on bacterial adhesion process and techniques used in studying bacterial-material interactions.  Biomatter. 2012;2(4):176-194.PubMedArticle
5.
Zoubos  AB, Galanakos  SP, Soucacos  PN.  Orthopedics and biofilm--what do we know? a review.  Med Sci Monit. 2012;18(6):RA89-RA96.PubMedArticle
6.
Renner  LD, Weibel  DB.  Physicochemical regulation of biofilm formation.  MRS Bull. 2011;36(5):347-355.PubMedArticle
7.
Donlan  RM.  Biofilms: microbial life on surfaces.  Emerg Infect Dis. 2002;8(9):881-890.PubMedArticle
8.
Fux  CA, Costerton  JW, Stewart  PS, Stoodley  P.  Survival strategies of infectious biofilms.  Trends Microbiol. 2005;13(1):34-40.PubMedArticle
9.
Branda  SS, Vik  S, Friedman  L, Kolter  R.  Biofilms: the matrix revisited.  Trends Microbiol. 2005;13(1):20-26.PubMedArticle
10.
Berbari  EF, Hanssen  AD, Duffy  MC,  et al.  Risk factors for prosthetic joint infection: case-control study.  Clin Infect Dis. 1998;27(5):1247-1254.PubMedArticle
11.
Karatan  E, Watnick  P.  Signals, regulatory networks, and materials that build and break bacterial biofilms.  Microbiol Mol Biol Rev. 2009;73(2):310-347.PubMedArticle
12.
Hoffman  LR, D’Argenio  DA, MacCoss  MJ, Zhang  Z, Jones  RA, Miller  SI.  Aminoglycoside antibiotics induce bacterial biofilm formation.  Nature. 2005;436(7054):1171-1175.PubMedArticle
13.
Anwar  H, Strap  JL, Chen  K, Costerton  JW.  Dynamic interactions of biofilms of mucoid Pseudomonas aeruginosa with tobramycin and piperacillin.  Antimicrob Agents Chemother. 1992;36(6):1208-1214.PubMedArticle
14.
Connaughton  A, Childs  A, Dylewski  S, Sabesan  VJ.  Biofilm disrupting technology for orthopedic implants: what’s on the horizon?  Front Med (Lausanne). 2014;1:22.PubMed
15.
Gallaher  TK, Wu  S, Webster  P, Aguilera  R.  Identification of biofilm proteins in non-typeable Haemophilus influenzae [published correction appears in BMC Microbiol. 2013;13:261].  BMC Microbiol. 2006;6:65.PubMedArticle
16.
North  JF.  The use of preserved bovine cartilage in plastic surgery.  Plast Reconstr Surg (1946). 1953;11(4):261-274.PubMedArticle
17.
Falcone  CL, Ogren  FP, Moore  GF, Yonkers  AJ.  Implants in nasal surgery.  Ear Nose Throat J. 1986;65(11):517-521.PubMed
18.
Romo  T  III, Pearson  JM.  Nasal implants.  Facial Plast Surg Clin North Am. 2008;16(1):123-132, vi.PubMedArticle
19.
Carter  WW.  The value of bone and cartilage transplants in rhinological surgery.  Ann Surg. 1917;66(2):162-168.PubMedArticle
20.
Metzenbaum  M.  Replacement of the lower end of the dislocated septal cartilage vs submucous resection of the dislocated end of the septal cartilage.  Arch Otolaryngol. 1929;9(3):282-296.Article
21.
Gillies  H.  A new graft applied to the reconstruction of the nostril.  Br J Surg. 1943;30:305-307.Article
22.
Laskin  DM, Sarnat  BG.  The metabolism of fresh, transplanted and preserved cartilage.  Surg Gynecol Obstet. 1953;96:493-499.
Original Investigation
Jul/Aug 2016

Analysis of Facial Implants for Bacterial Biofilm Formation Using Scanning Electron Microscopy

Author Affiliations
  • 1Department of Otolaryngology–Head and Neck Surgery, University of Illinois at Chicago
  • 2currently in private practice, Atlanta, Georgia
JAMA Facial Plast Surg. 2016;18(4):299-304. doi:10.1001/jamafacial.2016.0279
Abstract

Importance  Alloplastic implants are widely used in facial plastic surgery, both in rhinoplasty and nonrhinoplasty procedures. Implant infection and extrusion are significant concerns of such implants after placement. Bacterial biofilms have been previously implicated in chronic wound infections, particularly in the presence of foreign bodies, such as alloplastic facial implants. Owing to their structural composition, biofilms are resistant to treatment with conventional antibiotics, and implant removal is frequently the only option.

Objective  To evaluate explanted alloplastic facial implants for the presence or absence of bacterial biofilm using scanning electron microscopy.

Design, Setting, and Participants  Facial implants explanted by a single surgeon were analyzed for biofilm formation between July 1, 2012, and June 30, 2013. Of 7 facial implants, 4 consisted of silicone, and 3 were porous polyethylene. Six of the 7 were nasal dorsal implants, and 1 silicone implant was removed from the midface. Nonexplanted fresh silicone and porous polyethylene implants were each used as a control.

Main Outcomes and Measures  Scanning electron microscopy images were analyzed by an electron microscopist masked to the clinical history and implant type. The presence of biofilm formation was graded as none, mild, moderate, or severe.

Results  A total of 7 patients with previously placed alloplastic facial implants at an outside institution underwent revision rhinoplasty and removal of facial implants. All porous polyethylene implants showed biofilm formation to various degrees. Furthermore, all porous polyethylene implants had at least some areas of severe biofilm formation. One of the 3 porous polyethylene implants demonstrated severe biofilm formation on the entire implant, and the other 2 porous polyethylene implants showed areas of mild and severe biofilm formation. The only 2 implants without any evidence of biofilm were silicone implants. Of the other 2 silicone implants, 1 demonstrated no biofilm formation in 1 area and severe biofilm formation in another area, whereas the other had areas of moderate and severe biofilm formation.

Conclusions and Relevance  Five of 7 explanted facial implants showed at least some degree of biofilm formation. All implants with rougher surfaces, namely, porous polyethylene implants, demonstrated biofilm formation to a severe degree. Those with smoother surfaces, namely, silicone implants, were the only ones on which biofilm formation was either absent or less severe. Therefore, the suspicion that biofilms can form on facial implants is established through this investigation.

Level of Evidence  NA.

Introduction

Bacteria may exist in 2 different forms, namely, planktonic (free-floating) or biofilm.1,2 A biofilm is any group of microorganisms consisting of individual cells that stick to each other or to a surface, which could be either a living or nonliving surface.3 Although any surface is susceptible to biofilm growth, rougher and more hydrophobic surfaces will accumulate biofilms more rapidly.4 Once attached, the bacterial cells secrete an extracellular matrix consisting of DNA, proteins, and polysaccharides. The bacteria are ultimately embedded into this matrix to form a highly structured multicellular complex with numerous channels and cavities.2 In fact, biofilms may grow as much as 10 cm thick.5

Biofilm-forming bacteria have specialized structures, such as pili, fimbriae, flagella, and glycocalyx, which allow the cells to attach to an alloplastic implant.6 Beyond these specialized structures, adherence is further promoted by nonspecific factors, such as surface tension, hydrophobicity, and electrostatic forces.7 Once formed, biofilms are difficult to eradicate. Investigations have shown that chronic infections may be associated with bacteria living in biofilms, which makes such bacteria resistant to treatment with conventional antibiotics.8 This resistance is attributed, at least in part, to the fact that the structural composition of biofilms with their extracellular matrix does not allow for antibiotic penetration into such a highly organized arrangement.9

Biofilms on alloplastic implants may be monomicrobial or polymicrobial. Common organisms are Staphylococcus aureus, coagulase-negative staphylococci, β-hemolytic streptococci, and aerobic gram-negative rods (including Pseudomonas aeruginosa).10 Bacteria in biofilms are physiologically different from their free-floating counterparts. The differentiation into biofilm-forming bacteria is triggered by external factors, such as surface attachment, nutritional causes, or exposure to sublethal concentrations of antibiotics.11,12 The depletion of nutrients within biofilms leads to a slow-growing stationary state of the bacteria, which makes them up to 1000 times more resistant to antimicrobial agents than their free-floating counterparts.5 Therefore, eradicating biofilms with antibiotic therapy alone becomes a difficult task. Moreover, time is critical when treating presumed biofilm infections. During the first week, a biofilm may still be susceptible to certain antibiotics, such as tobramycin or piperacillin. Later on, the biofilm becomes resistant to antimicrobial treatment.13

Bacterial biofilm formation is a well-known problem in orthopedic surgery and ophthalmology. In recent years, biofilms have gained more interest in otolaryngology and facial plastic and reconstructive surgery as well. Any foreign material introduced into the human body predisposes these implants to infection. Treatment of such infections has become complicated because the alloplastic implants serve as a surface for bacterial colonization. Biofilm formation provides a protective milieu for the bacterial colonies on the implant, making them more resistant and difficult to eradicate when using standard antibiotic treatment. In some cases, the use of antibiotics alone has made the bacteria even more resistant to eradication. Therefore, there is a strong interest in developing antibiofilm, nonantibiotic agents to help disrupt biofilms on alloplastic implants.14

In the past, bacterial studies have focused on free-living (planktonic) bacteria, without investigating the highly structured and complex living arrangements of biofilms. Hence, it is critical to understand the molecular and microscopic structure of biofilms to develop adequate strategies to combat chronic biofilm infections.15 This study evaluated explanted alloplastic facial implants for the presence or absence of bacterial biofilms using scanning electron microscopy (SEM).

Box Section Ref ID

Key Points

  • Question Do various facial implant materials have different propensities to bacterial biofilm formation?

  • Findings Among a case series of 7 explanted alloplastic facial implants, all porous polyethylene implants showed biofilm formation to various degrees and had at least some areas of severe biofilm formation. The only 2 implants without any evidence of biofilm were silicone implants.

  • Meaning Alloplastic implants with rougher surfaces, such as porous polyethylene, appear to form biofilms more readily and more densely than implants with smoother surfaces, such as silicone.

Methods

This study was approved by the University of Illinois at Chicago Institutional Review Board. No informed consent for study participation was obtained from the patients because the implants were explanted with their permission.

Surgical Removal of Facial Implants

A total of 7 patients with previously placed alloplastic facial implants at an outside institution underwent revision rhinoplasty and removal of facial implants by one of us (D.M.T.) between July 1, 2012, and June 30, 2013. The alloplastic facial implants, either porous polyethylene or silicone, were surgically removed either owing to suspected infection or during revision surgery for other indications. A small piece of the implant measuring approximately 2 × 4 × 5 mm was sterilely cut from each implant’s surface in the operating room, placed in a sterile specimen cup, and kept moist with 0.9% normal saline. On completion of surgery, the specimen was immediately placed into a 3% gluteraldehyde with cacodylate buffer (0.1M) (Electron Microscopy Sciences) and refrigerated at 4°C.

Specimen Preparation and Imaging

The segments of explanted alloplastic implants were delivered refrigerated to the SEM laboratory at the University of Illinois at Chicago. The specimens were prepared for SEM analysis by a senior scanning electron microscopist. First, the specimens were dehydrated with a series of ethanol concentrations of 50%, 70%, 85%, 95%, and 100%. The specimens were subsequently immersed in hexamethyldisilazane for 15 minutes, air-dried, and mounted on aluminum studs (M4 tapered hole, female thread, 15-mm head, and 6-mm pin height; Hitachi S-450 M4) with double-sided carbon-coated tabs (both from Electron Microscopy Sciences). They were then sputter-coated with 4 nm of platinum and palladium using a high-resolution coater (208 HR; Cressington Scientific Instruments, Inc) before viewing the samples with a variable-pressure scanning electron microscope (V-P mode, S-3000N; Hitachi). Digital images were subsequently captured using PCI software.

Statistical Analysis and Evaluation

The captured SEM images of the 7 specimens were sent digitally to the scanning electron microscopist (masked to the clinical history and implant type) for analysis. These images were examined for the presence of biofilm formation and graded as none, mild, moderate, or severe (Figure). Nonexplanted fresh silicone and porous polyethylene implants were each used as a control.

Results

Of the 7 patients with facial implants, 6 had nasal dorsal implants, and 1 had a midface cheek implant. Four of the 7 implants consisted of silicone, and 3 were porous polyethylene. All 3 porous polyethylene implants were from the nasal dorsum, and 3 of the 4 silicone implants were from the nasal dorsum, with the fourth from the midface (Table).

Of the 7 explanted implants, the only 2 without any evidence of biofilm formation were silicone implants, with 1 being a nasal dorsal implant and the other being a midface implant. Of the other 2 silicone implants, 1 demonstrated no biofilm formation in 1 area and severe biofilm formation in another area, whereas the other had areas of moderate and severe biofilm formation. All porous polyethylene implants showed biofilm formation to various degrees, with all of them having at least some areas of severe biofilm formation. One of the 3 porous polyethylene implants demonstrated severe biofilm formation on the entire implant, and the other 2 porous polyethylene implants showed areas of mild and severe biofilm formation (Table). Neither the nonexplanted fresh silicone control nor the porous polyethylene control demonstrated any biofilm formation.

Discussion

In 1953, North16 outlined the following characteristics of an ideal surgical implant: (1) easily harvested without a significant or painful operation at the donor site; (2) well tolerated at the recipient site; (3) no tendency of perforating through the skin or mucosa when placed in proximity to those sites, even in areas that are subject to frequent trauma, such as the nose or ears; (4) minimal distortion tendencies; and (5) minimal risk of resorption. Beyond these criteria, infection is a significant problem that predisposes implants to resorption, extrusion, mobility, and other problems.

Rhinoplasties, among other facial operations, are not sterile procedures but rather clean-contaminated surgical procedures, increasing the risk of surgical site infections through introduction of bacteria at the time of the operation. Owing to the nature of these procedures, implant colonization with bacteria is a higher risk in clean-contaminated operations than in sterile procedures, such as joint replacements in orthopedic surgery, in which biofilms have been well studied.

Some of the earliest graft materials used in rhinoplasty were heterografts, consisting most often of either bovine bone or cartilage. However, these grafts triggered an intense inflammatory response at the recipient site and have hence been abandoned owing to the high rate of subsequent graft resorption.17,18

Autologous grafts quickly became preferred options, which may be either cartilaginous or bony in nature. Autologous cartilage grafts may be obtained from the nasal septum, the ear (concha) for smaller defects, or the rib for larger defects.1719 The first use of septal cartilage was reported by Metzenbaum20 in 1929, and grafting of composite auricular cartilage was described by Gillies21 in 1943. To date, many reports exist describing a high success rate of autologous cartilage grafts owing to their availability, unique structural composition, low metabolic demand, and relative avascularity. However, there are also disadvantages associated with such autologous cartilage grafts, such as donor site morbidity and relative limited availability.22

In light of these factors and despite certain disadvantages, autografts are often the preferred grafting material in facial plastic surgery, particularly in rhinoplasty. When available, autografts cause much less of an inflammatory response than heterografts or alloplastic implants, leading to a lower rate of tissue reaction, graft resorption, and possible wound infection.

However, alloplastic facial implants are widely popular. They are abundantly available, without any donor site morbidity. Surgical time is decreased, leading to potential lower surgical cost and possibly increased patient safety. Many alloplastic implants are also easy to sculpt, such as silicone or porous polyethylene. In facial surgery, alloplastic implants are commonly used for chin augmentation, midface augmentation, orbital reconstruction, and nasal dorsal augmentation. There is a wide range of implant materials, including titanium, silicone, porous polyethylene, and polytetrafluoroethylene, to name a few. Particularly in rhinoplasty, a concern regarding implant infection and extrusion exists with alloplastic implants. This consideration stems from the thin nasal skin and mucosal coverage of the implant, its dual-surface exposure, the clean-contaminated nature of the procedure rather than truly sterile surgery, and possible compromise due to subsequent nasal trauma. While some implants, such as silicone, are soft and easy to sculpt and do not trigger much of an inflammatory reaction, they may also have a higher risk of mobility given their lack of tissue incorporation and formation of a capsule. On the other hand, porous polyethylene exhibits tissue ingrowth and is more readily incorporated into the nasal skeletal framework, with a lower risk of implant mobility, but at the expense of a more intense fibrotic reaction. This limitation can ultimately lead to overlying skin changes, which may be permanent. The tight adherence and scar formation of porous polyethylene make implant removal a difficult task, if needed. The most feared complication with alloplastic implants is implant infection, with subsequent extrusion, either through the external skin or the internal mucosal lining. Alloplastic implants are at a higher risk for infection owing to (1) the lack of vascular ingrowth and hence decreased ability to fight an infection, if present, and (2) their propensity for biofilm formation. As outlined above, biofilms adhere to surfaces tightly and are difficult to eradicate, once present, owing to their biochemical and structural composition.

This study examined explanted alloplastic facial implants for biofilm formation. Of the 7 specimens analyzed, 5 showed biofilm formation to varying degrees. However, only 1 patient had an infection observed clinically, which was the indication for explantation of the implant. The remaining 6 patients had their implants removed for other clinical or aesthetic reasons, without any clinical signs of infection at the time of surgery.

Although biofilms can theoretically form on any surface, living or nonliving, it has been well established that biofilms form more readily on surfaces that are rough and hydrophobic.3,4 Furthermore, an infection is more difficult to eradicate in environments of relative avascularity. Alloplastic facial implants meet the characteristics of having rough surfaces and being hydrophobic. They are also avascular. Hence, it seems logical that biofilms can readily form on alloplastic facial implants, especially in the context of facial surgery, which is not truly sterile, and bacteria may likely be seeded at the time of implant placement. Furthermore, because alloplastic implants are more avascular than their autograft counterparts, an infection (once present) would also be more difficult to eradicate owing to the diminished blood supply. Biofilms have an avascular core and are hence the most resistant form of any bacterial infection state. Therefore, a biofilm infection on an alloplastic implant is likely the most difficult infection to treat.

Questions arise. What constitutes a clinical infection and what constitutes a colonization state only? Why do some patients with biofilms have no clinically relevant complications, while others have extrusion of the implant? What are the triggers for a biofilm to transform from a clinically insignificant colonization state into a pathologic infection? Although this study does not investigate these questions, we now know that silicone and porous polyethylene, the 2 alloplastic facial implant materials analyzed in this study, are able to promote and harbor biofilm formation.

When a patient sustains facial trauma with an infection in proximity to the implant (eg, a sinus infection) or an infection at a more distant site, the bacteria of such an infection may interact with the already present biofilm. This scenario in turn may then trigger the biofilm to transform from a colonization state into a clinical infection either via direct bacterial interaction at the biofilm surface or via an inflammatory signaling cascade. Irrespective of the exact underlying mechanism, some initial insult must exist that triggers such a transformation to occur to account for the fact that some alloplastic implants are only colonized with biofilms, without a clinically evident infection.

The limitations of this study are its small sample size (n = 7), lacking statistical power. Furthermore, quantification of biofilm formation is difficult and somewhat subjective in this study, although it was performed by an experienced scanning electron microscopist. True objective quantification of biofilm formation is challenging given the fact that biofilm formation on an implant’s surface is nonhomogeneous. Furthermore, only representative samples of a given implant rather than the entire implant were analyzed for technical reasons. Ideally, biofilm formation would be quantified and graded more objectively by measuring either biofilm density or thickness.

The surgical and postsurgical protocol at the time of implant placement is also variable given that different surgeons performed the initial placement of the implants. It is unknown if antibiotic irrigations were performed at the time of implant placement or if the patients received postoperative antibiotic prophylaxis. Most patients did not recall the exact date of alloplastic implant placement. In fact, 2 of the 7 patients were unaware of having an alloplastic implant altogether. Hence, no conclusion can be drawn if it is the implant type, location, time, surgical technique, or perioperative antibiotic protocol that contributes to biofilm formation. Furthermore, although present, biofilm formation may only be an incidental finding and clinically insignificant. Only 1 of the 7 patients (patient 4 in the Table) had chronic inflammation with overlying skin changes at the time of presentation for which secondary surgery was indicated. No acute infection or purulence was noted at the time of surgery.

Although we included a sterile porous polyethylene control and a silicone facial implant control that was not explanted from a patient, there is no true control group in this study. Furthermore, no alloplastic implants were compared with septal, ear, autologous rib, or irradiated rib grafts. Hence, no conclusion can be drawn about whether or not alloplastic implants are indeed more susceptible to infection than cartilaginous autografts.

However, despite these aforementioned limitations, this initial qualitative analysis confirms the suspicion that alloplastic facial implants are prone to biofilm formation. The fact that porous polyethylene implants appear to form biofilm more readily than silicone implants should also not come as a surprise given the rough and porous nature of polyethylene and the propensity of biofilms to form more readily on such surfaces. Further studies with a larger sample size, various alloplastic implant materials, and autografts are needed to compare different implant materials with respect to the tendency of biofilm formation and to draw a statistically significant conclusion.

Conclusions

This qualitative study illustrates that bacterial biofilms are present on explanted alloplastic facial implants. Furthermore, the results of this study suggest that alloplastic implants with rougher surfaces, such as porous polyethylene, form biofilms more readily and more densely than smoother implants, such as silicone. Scanning electron microscopy is an excellent tool to identify areas of biofilm formation. Further studies with a larger sample size are needed for a more objective analysis and quantification of biofilm formation, as well as to obtain statistical significance. No conclusion regarding alloplastic implants vs autografts with respect to their propensities of biofilm formation can be drawn from this initial analysis.

Back to top
Article Information

Accepted for Publication: March 7, 2016.

Corresponding Author: Thomas J. Walker, MD, 5555 Peachtree Dunwoody Road, Ste 135, Atlanta, GA 30342 (twalker@atlantaent.com).

Published Online: May 5, 2016. doi:10.1001/jamafacial.2016.0279.

Author Contributions: Both authors had full access to all the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.

Study concept and design: Both authors.

Acquisition, analysis, or interpretation of data: Walker.

Drafting of the manuscript: Walker.

Critical revision of the manuscript for important intellectual content: Both authors.

Statistical analysis: Walker.

Obtained funding: Toriumi.

Administrative, technical, or material support: Walker.

Study supervision: Both authors.

Conflict of Interest Disclosures: None reported.

Additional Contributions: Scanning electron microscopy was performed by Linda Juarez at the Electron Microscopy Service Laboratory at the University of Illinois at Chicago. Image analysis was performed by Patricia Blackwelder, PhD, at the Center of Advanced Microscopy at the University of Miami, Miami, Florida. Neither received compensation.

References
1.
Costerton  JW, Stewart  PS, Greenberg  EP.  Bacterial biofilms: a common cause of persistent infections.  Science. 1999;284(5418):1318-1322.PubMedArticle
2.
Stoodley  P, Sauer  K, Davies  DG, Costerton  JW.  Biofilms as complex differentiated communities.  Annu Rev Microbiol. 2002;56:187-209.PubMedArticle
3.
Hall-Stoodley  L, Costerton  JW, Stoodley  P.  Bacterial biofilms: from the natural environment to infectious diseases.  Nat Rev Microbiol. 2004;2(2):95-108.PubMedArticle
4.
Ribeiro  M, Monteiro  FJ, Ferraz  MP.  Infection of orthopedic implants with emphasis on bacterial adhesion process and techniques used in studying bacterial-material interactions.  Biomatter. 2012;2(4):176-194.PubMedArticle
5.
Zoubos  AB, Galanakos  SP, Soucacos  PN.  Orthopedics and biofilm--what do we know? a review.  Med Sci Monit. 2012;18(6):RA89-RA96.PubMedArticle
6.
Renner  LD, Weibel  DB.  Physicochemical regulation of biofilm formation.  MRS Bull. 2011;36(5):347-355.PubMedArticle
7.
Donlan  RM.  Biofilms: microbial life on surfaces.  Emerg Infect Dis. 2002;8(9):881-890.PubMedArticle
8.
Fux  CA, Costerton  JW, Stewart  PS, Stoodley  P.  Survival strategies of infectious biofilms.  Trends Microbiol. 2005;13(1):34-40.PubMedArticle
9.
Branda  SS, Vik  S, Friedman  L, Kolter  R.  Biofilms: the matrix revisited.  Trends Microbiol. 2005;13(1):20-26.PubMedArticle
10.
Berbari  EF, Hanssen  AD, Duffy  MC,  et al.  Risk factors for prosthetic joint infection: case-control study.  Clin Infect Dis. 1998;27(5):1247-1254.PubMedArticle
11.
Karatan  E, Watnick  P.  Signals, regulatory networks, and materials that build and break bacterial biofilms.  Microbiol Mol Biol Rev. 2009;73(2):310-347.PubMedArticle
12.
Hoffman  LR, D’Argenio  DA, MacCoss  MJ, Zhang  Z, Jones  RA, Miller  SI.  Aminoglycoside antibiotics induce bacterial biofilm formation.  Nature. 2005;436(7054):1171-1175.PubMedArticle
13.
Anwar  H, Strap  JL, Chen  K, Costerton  JW.  Dynamic interactions of biofilms of mucoid Pseudomonas aeruginosa with tobramycin and piperacillin.  Antimicrob Agents Chemother. 1992;36(6):1208-1214.PubMedArticle
14.
Connaughton  A, Childs  A, Dylewski  S, Sabesan  VJ.  Biofilm disrupting technology for orthopedic implants: what’s on the horizon?  Front Med (Lausanne). 2014;1:22.PubMed
15.
Gallaher  TK, Wu  S, Webster  P, Aguilera  R.  Identification of biofilm proteins in non-typeable Haemophilus influenzae [published correction appears in BMC Microbiol. 2013;13:261].  BMC Microbiol. 2006;6:65.PubMedArticle
16.
North  JF.  The use of preserved bovine cartilage in plastic surgery.  Plast Reconstr Surg (1946). 1953;11(4):261-274.PubMedArticle
17.
Falcone  CL, Ogren  FP, Moore  GF, Yonkers  AJ.  Implants in nasal surgery.  Ear Nose Throat J. 1986;65(11):517-521.PubMed
18.
Romo  T  III, Pearson  JM.  Nasal implants.  Facial Plast Surg Clin North Am. 2008;16(1):123-132, vi.PubMedArticle
19.
Carter  WW.  The value of bone and cartilage transplants in rhinological surgery.  Ann Surg. 1917;66(2):162-168.PubMedArticle
20.
Metzenbaum  M.  Replacement of the lower end of the dislocated septal cartilage vs submucous resection of the dislocated end of the septal cartilage.  Arch Otolaryngol. 1929;9(3):282-296.Article
21.
Gillies  H.  A new graft applied to the reconstruction of the nostril.  Br J Surg. 1943;30:305-307.Article
22.
Laskin  DM, Sarnat  BG.  The metabolism of fresh, transplanted and preserved cartilage.  Surg Gynecol Obstet. 1953;96:493-499.
×