Optometry & Vision Science:
The Impact of Contact Angle on the Biocompatibility of Biomaterials
Menzies, Kara L.*; Jones, Lyndon†
†PhD, FCOptom, FAAO
School of Optometry, University of Waterloo, Waterloo, Ontario, Canada.
Received August 24, 2009; accepted January 28, 2010.
Biomaterials may be defined as artificial materials that can mimic, store, or come into close contact with living biological cells or fluids and are becoming increasingly popular in the medical, biomedical, optometric, dental, and pharmaceutical industries. Within the ophthalmic industry, the best example of a biomaterial is a contact lens, which is worn by ∼125 million people worldwide. For biomaterials to be biocompatible, they cannot illicit any type of unfavorable response when exposed to the tissue they contact. A characteristic that significantly influences this response is that related to surface wettability, which is often determined by measuring the contact angle of the material. This article reviews the impact of contact angle on the biocompatibility of tissue engineering substrates, blood-contacting devices, dental implants, intraocular lenses, and contact lens materials.
Biomaterials are compounds of natural or artificial origin that can mimic, store, or come into close contact with living biological cells or fluids.1 Use of biomaterial has shown steady growth over the past half century, particularly in the areas of medicine, biology, materials science, and engineering. Specific examples of biomaterials include tissue engineering substrates, blood-contacting medical devices, artificial joint replacements, dental impressions, breast implants, ocular implants, and contact lenses. Table 1 includes approximate numbers of these devices used in the United States.2–6 These biomaterials are primarily used for medical applications; however, biomaterials can also be used for fertility regulation implants in cattle and cell-silicon biochips.7
Biomaterials are rarely used as isolated or independent materials for medical applications, being typically integrated into devices or implants. The integrated biomaterials are typically classified as being inert, active, or degradable, depending on their specified use. An inert biomaterial will illicit no or minimal tissue response, “active” biomaterials will encourage growth or attachment of surrounding tissue (e.g., a tissue graft), and degradable biomaterials typically dissolve over time (e.g., dissolvable sutures). Table 2 lists a variety of medical applications and the biomaterial used for each application.6,8–11
There are a number of factors that need careful consideration when designing a biomaterial, including toxicology, biocompatibility, healing, and dependence on specific anatomical sites of implantation, to name a few.7 One factor relates to the exposure time of the surrounding biological cells or fluids to the biomaterial. For example, a hemodialysis membrane may be in contact with a patient's blood for 3 hours, a contact lens may be worn for a couple of days or months, and a hip or joint replacement, intraocular lens (IOL), or heart valve are designed to be in a patient's body for life. The interaction between the biological cells or fluids with the biomaterial clearly should not illicit an unfavorable response, which is particularly relevant for those implants that will be in place for extended periods of time.
The performance of a biomaterial in situ is generally evaluated by its “biocompatibility,” which refers to the measured success of the interaction between the biomaterial and the biological cells for a specific biomedical task.1,12 If the biomaterial retards or affects the natural biological process for which it is intended to assist, the biomaterial would be considered incompatible.13 For example, in tissue engineering, if a substratum is used, which does not promote the growth of a smooth monolayer of cells on its surface, it would not be deemed biocompatible, because its “desired outcome” is to allow the growth of a tissue on its surface.
The surface of the biomaterial is the first component of the implant that comes into contact with the biological cells or fluids. Thus, biocompatibility will be influenced primarily by the surface characteristics of the biomaterial, particularly the wettability, surface chemistry of the exposed atoms, surface energy, and the surface topography.14,15
The focus of this article is to review whether a biomaterial with a high degree of wettability will be more biocompatible than a biomaterial with a low degree of wettability. Special reference will be made to blood-contacting devices, tissue-engineering substrates, dental impressions, IOL materials, and hydrogel contact lenses.
The term “wettability” refers to the ease with which a fluid spreads across a solid surface or more specifically how the fluid adheres to the solid surface.16–18 Wettability of a solid substrate is influenced by three forces: the surface tension of the solid, the surface tension of the liquid, and the interfacial tension. At the interior of the liquid, there are more neighboring molecules, when compared with that seen at the liquid surface. The attractive forces between the neighboring molecules in the interior region generate a lower energy state compared with the higher energy state at the surface. Thus, molecules try to remain in the bulk of the liquid, which creates a tendency for liquids to maintain a minimum surface area by remaining as droplets on a hydrophobic surface.19 The surface tension of the solid is similar to that of a liquid; however, the intermolecular bonding of molecules in the solid is tighter, and the inward pull is not enough to change the shape of the solid. A solid with a high surface tension acts to pull the liquid over the surface of the dry solid in an attempt to reduce the surface tension.20
The surface tension of the solid is countered by a force at the solid-liquid interface, which pulls the liquid away from the surface of the dry solid. This force is known as the “interfacial tension.” The force of the interfacial tension can be increased or lessened depending on the attractive forces between the molecules in the liquid and the solid. The greater the attraction between the liquid and the solid, the lower the interfacial tension will be and the more spreading of the liquid over the surface of the solid.
In the contact lens literature, wettability in-eye is typically assessed by determining the ease with which the tear film spreads on the contact lens surface and how stable the tear film remains adherent to that surface. This is usually achieved by visible inspection of the lens at the slit lamp,21 measuring the noninvasive tear breakup time,22–25 or determining the tear film thickness and stability using interference fringes.24,26,27 However, in the context of general biomedical materials, the surface wettability is usually assessed by determining water contact angles (CAs) at the material surface.
Measuring wettability of biomaterials in vitro is evaluated by measuring the CA at the liquid-solid interface. CAs are usually calculated using the Young-Dupré equation: cosθ = (γSV − γSL)/γLV, where γ is the interfacial tension between the solid (S), liquid (L), and vapor (V) phases.28 A high CA or high adhesion of the fluid to the solid indicates low wettability or a hydrophobic solid surface (Fig. 1a).29 A low CA, in which there is a smooth, continuous fluid film over the solid surface, signifies high wettability or a hydrophilic surface (Fig. 1b).
There are two different CAs that can be measured: advancing CA and receding CA. The advancing CA is the angle at which the liquid spreads across the surface of the dry solid at first contact. The receding CA is measured when the drop of liquid is being withdrawn from the surface of the solid. The receding angle is usually smaller than the advancing angle, because it is measuring a liquid being withdrawn over a surface that is already moist. The difference between the advancing and receding CAs is known as the “hysteresis.”16 Although hysteresis occurs because of withdrawing the droplet over an already wet surface, it is also thought to be caused by polymer reorientation at the material surface.30 Polymeric surfaces are mobile, and the orientation of the molecules can change depending on the surrounding environment. When the materials are exposed to air or other hydrophobic environments, the hydrophobic groups within the polymers will migrate toward the surface of the material, making the surface less wettable. For example, when poly(2-hydroxyethyl methacrylate) (pHEMA) is exposed to air, the methyl groups rotate toward the hydrophobic interface by chain rotation.30 This is a more favorable energetic state, thereby lowering the surface free energy. However, on exposure to polar liquids, the polymers will rotate so that the hydrophilic groups are pointing toward the polar phase. This increases the wettability of the solid surface.
Techniques for Measuring Wettability
There are three major techniques for measuring in vitro wettability: sessile drop, captive bubble, and the Wilhelmy balance method,20 with sessile drop being the most commonly used technique.31–38 The sessile drop technique involves placing a drop of liquid from a syringe onto the surface of the test material. After the drop is placed on the material, the advancing CA can be measured, usually with a goniometer. A goniometer captures an image of the drop on the test material and, through data analyses, the advancing angle at the liquid-solid interface is measured (Fig. 2).
The receding angle is measured by withdrawing the drop of liquid off the test material surface with a syringe, an image is taken as the drop is withdrawn, and the angle is measured between the liquid and the solid surface.33,39 Although the sessile drop method is widely used in measuring wettability, there are two major problems with the method, namely evaporation of the liquid and dehydration of the solid surface. A small drop has a large surface area compared with the volume of the liquid, and evaporation will cause the drop to retreat on the surface of the solid and affect the measured CAs.20 Dehydration of the solid surface can also lead to altered CAs. As mentioned earlier, polymers may reorient themselves at the surface in respect to the surrounding environment. If the polymer is left in a dry environment (e.g., air), those hydrophobic groups within the polymer will reorient themselves to the surface of the material, making the material less wettable. Both of these problems can be solved by placing the solid sample and liquid drop in a vapor-tight chamber with clear windows so that the image can still be captured and the CAs measured.20
The captive bubble technique eliminates the problem of the solid surface becoming dehydrated, because in this technique, the solid material is immersed in the probe liquid. A capillary tube is placed underneath the solid, and an air bubble is dispensed from the capillary tip so that the bubble just touches the surface of the solid (Fig. 3).40 As the air bubble becomes larger, it pushes the liquid away from the solid surface, and at this point, the angle between the solid and liquid is the receding angle. The air bubble is then retracted back into the syringe, and the liquid spreads back onto the surface of the lens. This angle would be considered the advancing angle.33
The captive bubble technique also has some drawbacks. First, because the sample is immersed in liquid, it may retain an absorbed layer of fluid on its surface, making it appear more wettable than it truly is in the natural environment. Second, the air bubble is a medium of low refractive index but is observed in a medium of high refractive index. This creates a light path that is from air-to-water to air-to-water and then back to air. This path makes it difficult to see where the bubble actually contacts the solid surface.20,30 Third, the amount of probe liquid required makes the captive bubble technique relatively expensive to perform.
The Wilhelmy balance method uses a plate or sample of test medium, which is hung from a microbalance and slowly submerged and removed vertically from a test liquid (Fig. 4). The advancing CA is the angle between the solid and the meniscus of the test liquid because the solid is dropped into the liquid, and the receding angle is the angle between the solid and the liquid because the plate is moved out of the liquid. The main disadvantage of this technique is the cost of the equipment and the time and expertise required to prepare the test sample for immersion.41,42
The accuracy of all the three methods depends primarily on how efficiently the experimenter prepares the solid sample. A recent experiment determined the repeatability of CA measurements on contact lenses using the captive bubble and sessile drop techniques.43 The results showed that the CAs measured using the sessile drop technique were less repeatable than those measured using the captive bubble technique. This was thought to be primarily due to variability in blot drying the contact lens during the preparation of CA measurement using the sessile drop technique.43
All three methods may give different CA measurements for the same material. Shang et al.44 looked at the wettability of five solid colloids (smectite, kaolinite, illite, goethite, and hematite) using a variety of methods, including sessile drop and Wilhelmy plate methods. The largest difference in measured CAs was seen for hematite, with a higher CA measured using the Wilhelmy balance method. Hematite had the smoothest surface of the five solid colloids, suggesting that other properties of the solid colloids other than surface roughness influence surface wettability.
A recent study from our laboratory investigated the wettability of five silicone hydrogel (SH) materials, immediately after the lenses were removed from the blister pack and after a 48-hour soak in preservative free saline, to remove the influence of any surface-active agents in the blister-pack saline. Advancing CAs were measured using both the sessile drop and Wilhelmy balance methods (Table 3). The advancing CAs measured for asmofilcon A and balafilcon A lenses were not statistically different from each other for both techniques (p > 0.05). The advancing CA for galyfilcon A measured by the Wilhelmy balance method was statistically lower than the CA measured using the sessile drop technique (p < 0.05). However, the CAs measured using the Wilhelmy balance technique were statistically higher (p < 0.05) than the CAs measured using the sessile drop technique for the lotrafilcon B and comfilcon A materials. The lower CAs measured using the Wilhelmy balance are probably due to the way the lenses are prepared, in that the lenses are not blot dried but rather the excess fluid is merely shaken off the lens surface before immersion, resulting in a layer of fluid being retained at the material interface. Because of the obvious differences in CAs measured using the different techniques, it is clearly important to accurately describe the technique used when reporting CAs of different biomaterials.
Impact of Contact Angle on Biomaterial Compatibility
Much work has been undertaken on measuring and reporting the CAs obtained with a variety of biomaterials. It has long been assumed that enhanced wettability (as evidenced by a low CA) will result in improved biocompatibility.45,46 The remainder of this review will investigate whether this concept is borne out in fact, by examining the literature for tissue engineering substrates, blood-contacting devices, dental impressions, IOLs, and finally hydrogel and SH lens materials. In addition, the impact of CA on the development of biofilms is also reviewed because these can lead to substantial issues with biomaterials, ultimately resulting in failure of the device in question.
Tissue Engineering Substrates
Tissue engineering incorporates the combination of cells, engineering, and biochemical and physiochemical factors to repair or replace portions of (or whole) tissue,47 and it holds great promise in medicine for patients that undergo extreme tissue and organ damage. Engineered tissues consist of three components (a) donated stem or precursor cells, (b) scaf-fold substrates, and (c) growth factors that will promote or prevent cell adhesion, proliferation, and differentiation.48–50 Of these three components, the scaf-fold substrate has the biggest impact on the generation of a biocompatible biomaterial and thus is the main interest in tissue engineering. If the scaf-fold allows for the growth, migration, and differentiation of cells on its surface, it will be deemed biocompatible.47,48
Tissue engineering begins by seeding the anchorage-dependent cells on the artificial substrate, which then attach, grow, migrate, and proliferate to form a monolayer of cells.51 During this process, the cell-substrate interaction is of most importance and must dominate over any other cell-cell interactions.47 Cell adhesive serum proteins such as fibronectin, fibrinogen, and vitronectin facilitate the adhesion of cells to the biomaterial; hence, the biomaterial must also allow the adhesion of serum proteins.52 After the monolayer is formed, cell-cell interactions become the principle factor in controlling cell migration and proliferation in the developing tissue.53
Many successful studies have been undertaken using self-assembling monolayers (SAMs) to create materials with a desired molecular composition and improved surface properties. Faucheux et al.54 used SAMs of organosilanes to prepare surfaces with varying wettabilities to test the effect of wettability on cell adhesion. Advancing and receding CAs were measured in this study by the sessile drop technique, using ultrapure water as the probe liquid dispensed from the syringe.54 SAMs were created with varying wettabilities: hydrophobic (CA > 80°), moderately wettable (CA = 48–62°), and hydrophilic (CA < 35°). These surfaces were exposed to the cell adhesive serum proteins, fibronectin, and vitronectin. The results showed that adherence of the proteins was limited on the highly hydrophobic and highly hydrophilic surfaces. The biomaterials were then exposed to human fibroblast cells. The cells that adhered to the moderately wettable surfaces showed a linear growth of the cells, whereas on the “extremely” hydrophobic and hydrophilic surfaces, the growth of the fibroblasts was greatly reduced, probably due to the absence of adhesion proteins.54 This demonstrates that the adhesion of the cells was clearly mediated by the adhesion of the proteins, which are significantly influenced by the surface wettability of the material under examination.
In a more recent study by Arima and Iwata,55 three SAMs were created, all of which exhibited varying wettabilities, as determined using the sessile drop technique. The SAMs were then exposed to endothelial and epithelial cells. Results after the SAMs were exposed to the endothelial cells showed an increase in adhesion as the wettability increased. As in the previous study,54 maximum endothelial cell adhesion was seen at a CA of ∼35 to 50° depending on the surface chemistry of the SAM.55
Tzoneva et al.47 examined the adhesion of extracellular matrix (ECM) proteins, such as fibronectin, fibrinogen, and collagen, on two artificial substrates. One substrate was hydrophilic with an advancing CA of ∼24°, and the other substrate was hydrophobic with an advancing CA of ∼86°. Results showed that absorption of the proteins onto hydrophobic surfaces was less than that on hydrophilic surfaces, as expected. Those proteins that did adhere to the hydrophobic surface underwent a conformational change leading to a decreased accessibility of focal adhesion sites to allow the formation of the monolayer of endothelial cells. Absorption of the ECM proteins on the hydrophilic surface was much higher than that on the hydrophobic surface. However, when looking at cell-cell contact after the precoated hydrophilic surfaces were exposed to endothelial cells, there was an inability for the cells to interact with each other and form a smooth consistent monolayer. This appeared to be related to the stronger cell-substrate adhesion of the ECM proteins to the hydrophilic surface.
From these three experiments, it appears that a highly wettable scaf-fold for tissue engineering does not necessarily lead to a more biocompatible scaf-fold, with moderately wettable surfaces resulting in enhanced biocompatibility. More research is required to develop a material that has the ideal balance of hydrophobic and hydrophilic surfaces, such that there will be strong cell-substrate interaction and subsequent strong cell-cell interactions, to create the smooth monolayer required for tissue regeneration.
Biomaterials are frequently used as blood-contacting devices in modern medicine.8 These devices include stents, prosthetic heart valves, vascular grafts, hemodialysis membranes, vena cava filters, and blood bags.8 In the presence of a biomaterial, platelets tend to adhere to the foreign material and become activated. Once activated, the platelets initiate thrombosis by secreting prothrombotic factors leading to clotting.56 Under normal conditions, thrombosis and complement activation are favorable responses preventing blood loss. However, in the presence of a foreign material such as a blood-contacting device, thrombosis, and complement activation is unfavorable leading to blood clots and hence low biocompatibility.8 Thus, a blood-contacting device that does not elicit thrombosis would be considered biocompatible.
The surface properties, particularly the wettability of a biomaterial, are highly important when blood comes into contact with the biomaterial. Platelets tend to respond differently to hydrophobic or hydrophilic monomers.57 Therefore, when coating a biomaterial, the effect of wettability on the activation of platelets must be carefully examined.
Two recent studies have been undertaken in which nitrogen-doping on plasma-implanted silicone and amorphous carbon films was examined.58,59 The sample films under investigation in both studies varied in their degree of wettability as determined by measuring the CAs of the surface by the sessile drop technique. Results from both studies showed a decrease in platelet activation with an increase in wettability.58,59 The morphology of the cells that adhered to the hydrophobic surface showed spreading, accompanied by secretion and synthesis of multiple thrombotic factors.
Synthetic blood-contacting membranes are becoming increasingly popular for use in medical applications.60 These membranes are used in treatments such as hemodialysis, hemodiafiltration, hemofiltration, plasmapheresis, plasma collection, and oxygenation of blood during cardiac surgery.8,60 These membranes require carefully designed bulk properties for the desired permeability, adsorption, and blood compatibility for their specific uses.60 For treatments such as plasma collection or oxygenation, it is less crucial to have a high compatibility between blood and the membrane, because the treatment time is relatively short. However, in chronic dialysis, the treatment duration can be up to 3 to 5 hour.60 Long-term complications and renal disease may be attributed to low biocompatibility between blood and the synthetic membrane during dialysis.
Protein adsorption is a substantial problem in terms of compatibility between blood and the membrane.60 Studies have been conducted in which membranes with different wettabilities were created and tested. These studies found that a membrane with a balanced distribution of hydrophobic and hydrophilic domains had the best compatibility and led to a reduction in protein adsorption and a reduction in coagulation.60,61 Thus, for blood-contacting devices (as with tissue engineering substrates), the literature would suggest that biomaterials with a high degree of wettability and low CA do not necessarily result in enhanced biocompatibility.
Dental impressions are used for the fabrication of dentures, crowns, and models for people who require dental braces. A dental impression is created first by placing a viscous liquid material into a person's mouth around the teeth and gums,62 typically in a custom container. The liquid material is allowed to set, after which it is removed, leaving an impression of the person's teeth and gums. A gypsum material is poured into the impression and allowed to harden to produce a replica of the teeth and gums. Common materials used for impressions consist of polyether and silicone.63
The material used for the impressions is highly important, in that it needs to form readily around the teeth and gums and be void of any bubbles or defects. The environment inside the mouth is moist due to the excretion of saliva; and therefore, the material used for dental impressions must be hydrophilic to ensure compatibility with the environment inside the mouth.63 It is also important that the material is hydrophilic both before and after setting. The material should be hydrophilic before setting so that it can easily form around the teeth and gums and needs to remain hydrophilic after setting so that no air bubbles are entrapped when the gypsum products are poured. If a surface is hydrophobic, water, saliva, or any other fluid on the surface of the teeth or gums would create a small droplet and hence create a small void in the impression material or gypsum.64,65 This was confirmed by Michalakis et al.,63 who looked at the impact of wettability on void formation. The wettability of six materials was determined using the sessile drop method. The results showed that the material with the lowest CA, polyether, exhibited the fewest voids before setting. In addition, the materials with a CA of ≤60.8° exhibited fewer voids on the surface of the casted material. Thus, examination of the literature would suggest that a low CA does indeed result in a more biocompatible dental impression material.
Cataract surgery is the most common surgical procedure conducted, with >1.6 million cataract surgeries a year being undertaken in the United States alone.66 The general surgical method used consists of the removal of the crystalline lens and subsequent insertion of an IOL. Historically, IOLs were made of polymethylmethacrylate (PMMA), which were generally lathe cut from a PMMA rod or button.66–69 PMMA was originally regarded as being inert having a high biocompatibility in the eye; however, studies showed that PMMA caused postsurgical adhesion of inflammatory cells on the surface of the IOL,70–73 uveitis, breakdown of the blood-aqueous barrier, and potential loss of vision.10,66 These adverse reactions were believed to be related—at least in part—to the hydrophobic nature of PMMA, which led to the development of hydrophilic heparin surface-modified PMMA lenses.67,68 This material modification demonstrated decreased levels of cell adhesion to the surface of the IOL.68,74,75 However, with the invention of small incision cataract surgery, foldable IOLs were created. Foldable IOLs are currently made primarily from hydrophobic or hydrophilic acrylic or silicone-based materials.67,69,76
Much research has been undertaken investigating the biocompatibility of the different materials used to make IOLs and the impact of surface wettability. Biocompatibility of IOLs is most commonly based on the proliferation of anterior lens epithelial cells (LECs) onto the surface of the IOL, resulting in the development of anterior capsular opacification and posterior capsular opacification. In 2006, Yao et al.77 investigated in vitro the attachment and proliferation of LECs on the surface of a foldable silicone IOL and a surface modified foldable silicone IOL. The surface modified IOL had phospholipid-containing monomers on the surface of the IOL to create a more hydrophilic surface. The wettability of the silicone IOL and surface modified IOL was determined using the sessile drop technique. Both types of IOLs were exposed to LECs, and the results showed a significantly lower amount of LEC attachment and proliferation on the surface-modified IOL. Thus, the result of this study indicated that the more hydrophilic IOL had a greater biocompatibility than the more hydrophobic IOL.
Schroeder et al.78 investigated the adhesion of fibronectin onto the surface of IOLs with differing wettabilities. It is thought that attachment of fibronectin onto the surface of an IOL further enhances the binding of cellular and extracellular molecules, leading to complications such as posterior and anterior capsular opacifications.78 The wettability of the IOLs used in this study was determined using the sessile drop technique. The IOLs with lower CAs attracted the most fibronectin when compared with the IOLs with higher CAs. Thus, the results from this study contradicted those of Yao et al.,77 demonstrating that hydrophobic IOLs are more biocompatible than hydrophilic IOLs because of a lower amount of fibronectin absorbed onto the surface of the lens.
From the studies summarized earlier, it would appear that more research is required to determine if there is any correlation between CA values and IOL biocompatibility. It may transpire that the impact of CA on biocompatibility depends on the protein or cell-type under investigation.
Hydrogel Contact Lenses
There are a number of factors that influence the biocompatibility of contact lenses. The lens material must support a continuous tear film for optimum visual clarity, it must not dehydrate, and it needs to resist sorption of tear components such as lipids, proteins, and mucins, because buildup of deposition can lead to decreased visual clarity and reduced comfort.79 In addition, the cornea requires oxygen to maintain its clarity, structure, and function, thus, the contact lens material must be highly permeable to oxygen.66
Soft contact lenses were initially commercialized in 1970 and were composed of solely pHEMA.11 These lenses were rapidly accepted by patients and practitioners because of their relative comfort over PMMA lenses and adequate wettability.80 However, overnight wear of pHEMA-based lenses led to hypoxia and marked neovascularization.81 Despite many attempts over the next 30 years to enhance the oxygen transmissibility of pHEMA-based materials, through the addition of various monomers and changes in design, adequate oxygen transmissibility to support overnight wear was never possible, which led to the development of SH contact lenses.82
SH contact lenses became commercially available in 1999. They incorporated silicon as siloxane (—Si(CH3)2―○―) polymers into the lens material to improve oxygen permeability. Although siloxane is highly oxygen permeable, it is also highly hydrophobic, resulting in poorly wettable surfaces. For these materials to wet adequately in-eye, they require some form of modification to enhance their surface wettability. Initially, companies surface-treated the lenses by either plasma surface treatment or plasma oxidation.11,83,84 Table 4 shows a list of all SH contact lenses that are currently on the market.
Plasma is a highly reactive gas by activation of an electric field.13,84 When plasma is placed in the presence of a contact lens, the surface of the lens and the plasma react. Depending on the gases and compounds that are present during the plasma treatment process, different results will occur after the reaction has taken place. In the case of plasma oxidation, as used by Bausch & Lomb for the balafilcon A lens, oxygen is present in the plasma, which oxidizes the siloxane groups to silicate. This results in what resembles “islands of glass” on the surface of the lens,35,85–87 which are “clumps” of silicate. These silicate islands do not cover the entire surface of the lens and do not affect the oxygen permeability of the underlying balafilcon A material; however, their distribution is such that they increase the wettability of lens and thus support a stable tear film.82,85
The two CIBA Vision SH lenses, lotrafilcon A and lotrafilcon B, have a plasma coating in which volatile organic compounds are present in the plasma.11,35 When a reaction occurs, these compounds can behave as monomers and polymerize onto the surface of the lens, creating a polymer film that is more wettable than the underlying lens surface. This polymer layer is ultrathin (25 nm),88 does not affect the oxygen permeability of the underlying material, and results in a surface that is smoother than that measured on the balafilcon A lens.11,89
Johnson & Johnson do not use plasma treatments to improve the wettability of their SH lenses. Instead, they incorporate a high molecular weight wetting agent based on polyvinylpyrrolidone into the polymer matrix of the lens. Polyvinylpyrrolidone in aqueous environments readily binds to water retaining moisture for the lens. This treatment also helps to support a stable tear film and does not interfere with the oxygen permeability of the lens.90,91
Asmofilcon A (PremiO) is a relatively new SH material from Menicon. It uses a surface treatment known as Nanogloss, which combines plasma oxidation and plasma surface treatment, creating a smooth surface on the lens.92 To date, there is no published data on what monomers are incorporated into the lens material.
CooperVision manufactures two SH lenses, comfilcon A (Biofinity) and enfilcon A (AVAIRA). Neither of these lenses have any surface treatment or internal wetting agent. Rather, these lenses contain two silicone-based macromers that are incorporated into the lens material with hydrophilic monomers, resulting in a lens with a relatively high degree of wettability.93
Fig. 5, from our laboratory, displays the advancing CAs determined directly out of the blister pack and after soaking a variety of lenses for 48 hour to remove any potential impact of the packaging solution. The CAs measurements were carried out using the sessile drop technique. From Fig. 5, it can be seen that lenses with the plasma surface coating have a relatively low CA as determined by the sessile drop method and thus should exhibit a high degree of wettability.
Contact lenses that exhibit a high degree of in vitro wettability (as shown by having a low CA) should theoretically improve in-eye comfort.91 In a recent study, Cheung et al.94 compared a pHEMA-based hydrogel (etafilcon A) to that of a SH (galyfilcon A) in terms of comfort, ocular performance, and surface deposits. Participants wore the lens materials as a contralateral pair for 8 to 12 hour a day for 6 consecutive days. The results indicated that the lenses were comparable in comfort. However, our sessile drop CA measurements (Fig. 5) for galyfilcon A revealed a CA of 102°, compared with 51° for etafilcon A (internal data), suggesting that the etafilcon A material should be substantially more comfortable if CA was an important factor affecting lens comfort.
Keir et al.95 conducted a study to determine if there was any correlation between lens comfort and both in vivo wettability (determined via prelens noninvasive tear breakup time) and ex vivo wettability (measured using the sessile drop technique). Participants were assigned to wear lotrafilcon B and senofilcon A lenses contralaterally for 14 d. As with the study by Cheung et al.,94 the results showed no correlation between in-eye comfort and either in vivo or ex vivo wettability.95 Dumbleton et al.96 investigated comfort ratings throughout the day for five SH lenses (galyfilcon A, senofilcon A, lotrafilcon A, lotrafilcon B, and balafilcon A) with widely varying CAs (Fig. 5). The results indicated no significant difference between any of the lens types for comfort, burning, dryness, and overall comfort throughout the course of the day. In 2002, Morgan and Efron97 compared the comfort rating of balafilcon A (PureVision) and lotrafilcon A (Focus Night & Day). Subjects in this cross-over study wore a pair of each lens type for 8 weeks. Our sessile drop CA measurements (Fig. 5) for balafilcon A revealed a CA of 84°, compared with 42° for lotrafilcon A, suggesting that the lotrafilcon A material should be substantially more comfortable. However, the results showed that there was no difference in comfort rating between the two lens materials.97
The aforementioned four experiments suggest that there is a poor correlation between sessile drop CA and in-eye comfort, indicating that a low CA does not result in improved comfort or biocompatibility for contact lenses. Deposition of substances from the tear film such as proteins and lipids can cause reduced comfort and vision,98–100 and this deposition is influenced by material surface charge, wettability, water content, wear time, and tear film composition.101–103 To date, relatively little is known about the impact of this deposition on wettability, but two studies28,35 would suggest that wettability may initially be improved by small amounts of deposited tear components. Cheng et al.28 looked at the influence of lysozyme and mucin deposition on lens wettability by determining the captive bubble CAs of balafilcon A, lotrafilcon A, and etafilcon A. The lenses were exposed to saline, lysozyme, mucin, and a complex solution containing both mucin and lysozyme. The CAs of all three materials reduced significantly after being soaked in the lysozyme-containing solutions, suggesting that the protein deposition improved the lens wettability. Lorentz et al.35 investigated the impact of lipid deposition on the sessile drop CAs obtained with five SH materials (senofilcon A, balafilcon A, lotrafilcon A, lotrafilcon A, and galyfilcon A) and four conventional pHEMA-based materials (polymacon, omafilcon A, alphafilcon A, and etafilcon A). All materials were soaked in a PBS (phosphate-buffered saline) solution, low concentration lipid- containing tear solution (LTS), and high concentration LTS for 2 and 5 d, after which the CA was measured for each material. All conventional lenses, regardless of their initial wettability, had CAs of ∼35° after being soaked in a high concentration LTS for 5 d. The SH lenses with a plasma coating (lotrafilcon A and lotrafilcon B) showed a significant reduction in measured CA, particularly after being soaked in the high concentration LTS, which was attributed to lipid deposition onto the surface of the lens. From the earlier results, it would appear that a low CA, whichever method is used, does not necessarily improve the comfort or biocompatibility of contact lens materials.
The buildup of proteins or cells onto the surface of any of the biomaterials described earlier can impact on the subsequent binding of microbes to the underlying substrates. These microbes have the potential to cause adverse reactions in the body on initial infection or may colonize and form a “biofilm,” which may remain inert but later cause severe health issues.
Biofilms are “matrix-enclosed bacterial populations that are attached to each other and/or other surfaces, including microbial aggregates and floccules and also adherent populations within the pore spaces of porous materials.”104 The formation of a biofilm is initiated by a bacterial cell irreversibly binding to a surface using exopolysaccharide glycocalyx polymers.105 The following cell division leads to the formation of microcolonies bound within the glycocalyx matrix.105 Eventually, a biofilm will develop, consisting of single cells and microcolonies, all embedded in a matrix onto which recruited bacterial cells will adhere.104,105
Formation of a biofilm on a medical device can cause subsequent infection and failure of the medical device, particularly in immunocompromised patients.105–108 Consequently, bacteria in the form of a biofilm are less susceptible to antibiotics than planktonic (or free-floating) bacterial cells,105,109 which poses a significant problem when attempting to remove the biofilm. Increasing the dose of a systemic antibiotic to deal with the localized problem may be a solution; however, this may lead to the evolution of bacterial strains resistant to the antibiotic.Cells of the immune system that have been developed in response to the biofilm may cause more harm to the host as well. For example, antibodies produced in response to the antigens at the biofilm surface cannot penetrate the glycocalyx matrix and thus will form immune complexes, which often damage the surrounding tissue.105 Thus, surface properties of the biomaterial must be optimally designed to prevent the adhesion of unwanted bacterial cells.
Many studies have been undertaken investigating the influence of surface properties, such as wettability and surface roughness, on the adhesion of bacterial cells.110–113 There are conflicting results from studies examining the relationship between surface wettability and bacterial adhesion. Some studies report that bacterial adhesion is higher on hydrophobic surfaces,113–115 and other studies show the reverse, with highly hydrophilic substrates showing increased biofilm development.110,116 In a recent study by Tang et al.,110 five silicone samples with varying surface wettability and roughness were used to examine the corresponding changes in adhesion and colonization of Staphylococcus epidermidis.110 Surface wettability was determined using the sessile drop technique, with CAs ranging from ∼110 to 141°. The results showed that there was a decreasing order of bacterial adhesion with increasing hydrophobicity. However, when the bacterial adhesion on surfaces with similar hydrophobicities but differing surface roughness was examined, the material with the higher degree of surface roughness had a significant increase in bacteria adhesion. From this result, it was believed that surface roughness may have more influence on bacterial adhesion than surface hydrophobicity.110
The results by Tang et al.110 were not supported by those of Vermeltfoort et al.,117 who looked at the bacterial transfer from contact lenses to surfaces with different surface roughness and wettability. The sessile drop CAs for the materials used in this study ranged from 30 to 107°. This latter study showed that materials with the roughest surface and the highest CA supported the least amount of transferred bacteria. Bruinsma et al.116 investigated bacterial adhesion on hydrophilic and hydrophobic contact lenses. The wettability of the lenses investigated was determined using the sessile drop technique with the range of CAs measured being 57 to 106°. The adhesion of a gram-negative strain and a gram-positive strain of bacteria was tested, and the results indicated greater adhesion of both strains on the more hydrophilic contact lens material compared with the adhesion that occurred on the hydrophobic lens material. These conflicting results on the influence of surface wettability on bacterial adhesion indicates that there may be other underlying factors, such as surface roughness, that are more relevant in promoting or preventing the adhesion and colonization of bacterial cells, with CA playing a relatively minor role on this aspect of biocompatibility.
This review has substantiated that biomaterial surfaces exhibiting low CAs do not necessarily exhibit enhanced biocompatibility. Blood-contacting devices and tissue engineering substrates need an appropriate balance of hydrophilic and hydrophobic surface entities, because excessively hydrophobic surfaces enhance cell affinity and reduce biocompatibility, but highly hydrophilic surfaces prevent cell-cell interactions, which are particularly important in tissue engineering. Dental impressions need a low CA on the surface to have a high biocompatibility, whereas the wettability of IOLs appears to have an inconsistent effect on biocompatibility of the IOLs. Modern generation SH contact lenses use a variety of approaches to enhance their hydrophilicity and moderate their interaction with the ocular surface and tear film. Further developments to enhance their wettability could enable them to support a more stable tear film for a longer period of time, resulting in enhanced in-eye wettability and potential comfort improvements, but to date, laboratory-based CA analysis does not predict in-eye performance. Finally, CA analysis is poorly predictive of the development of bacterial biofilms, with other surface properties appearing to have greater relevance. Although data from in vitro evaluations of various biomaterials may aid in predicting some of their behavior in vivo, much work remains to develop more sophisticated methods to investigate biomaterial surface properties and to assist in predicting their long-term performance.
We thank Dr. Rachael Peterson for her help with the preparation of the article.
School of Optometry
University of Waterloo
200 University Avenue West,
ON N2L 3G1, Canada
1.Vogler EA. Structure and reactivity of water at biomaterial surfaces. Adv Colloid Interface Sci 1998;74:69–117.
2.Kodjikian L, Burillon C, Roques C, Pellon G, Renaud FN, Hartmann D, Freney J. Intraocular lenses, bacterial adhesion and endophthalmitis prevention: a review. Biomed Mater Eng 2004;14:395–409.
3.Barr JT. 2004 annual report. Contact Lens Spectrum 2005;20:26–31.
4.Kurtz S, Ong K, Lau E, Mowat F, Halpern M. Projections of primary and revision hip and knee arthroplasty in the United States from 2005 to 2030. J Bone Joint Surg Am 2007;89:780–5.
5.Brown SL, Middleton MS, Berg WA, Soo MS, Pennello G. Prevalence of rupture of silicone gel breast implants revealed on MR imaging in a population of women in Birmingham, Alabama. AJR Am J Roentgenol 2000;175:1057–64.
6.Ramakrishna S, Mayer J, Wintermantel E, Leong KW. Biomedical applications of polymer-composite materials: a review. Compos Sci Technol 2001;61:1189–224.
7.Ratner BD, Hoffman AS, Schoen FJ, Lemons JE. Biomaterials science: a multidisciplinary endeavor. In: Ratner BD, Hoffman AS, Schoen FJ, Lemons JE, eds. Biomaterials Science: An Introduction to Materials in Medicine, 2nd ed. Boston: Elsevier Academic Press; 2004:1–9.
8.Spijker HT, Graaff R, Boonstra PW, Busscher HJ, van Oeveren W. On the influence of flow conditions and wettability on blood material interactions. Biomaterials 2003;24:4717–27.
9.Lee CH, Singla A, Lee Y. Biomedical applications of collagen. Int J Pharm 2001;221:1–22.
10.Abela-Formanek C, Amon M, Schild G, Schauersberger J, Heinze G, Kruger A. Uveal and capsular biocompatibility of hydrophilic acrylic, hydrophobic acrylic, and silicone intraocular lenses. J Cataract Refract Surg 2002;28:50–61.
11.Nicolson PC, Vogt J. Soft contact lens polymers: an evolution. Biomaterials 2001;22:3273–83.
12.Black J. Biological Performance of Materials: Fundamentals of Biocompatibility, 3rd ed. New York: Marcel Dekker; 1999.
13.Chu PK, Chen JY, Wang LP, Huang N. Plasma-surface modification of biomaterials. Mater Sci Eng 2002;36:143–206.
14.Raffaini G, Ganazzoli F. Understanding the performance of biomaterials through molecular modeling: crossing the bridge between their intrinsic properties and the surface adsorption of proteins. Macromol Biosci 2007;7:552–66.
15.Lee JH, Khang G, Lee JW, Lee HB. Interaction of different types of cells on polymer surfaces with wettability gradient. J Colloid Interface Sci 1998;205:323–30.
16.Maldonado-Codina C, Efron N. Dynamic wettability of pHEMA-based hydrogel contact lenses. Ophthal Physiol Opt 2006;26:408–18.
17.French K. Contact lens material properties. Part 1: wettability. Optician 2005;230:20–28.
18.Johnson RE Jr, Dettre RH. Wetting of low-energy surfaces. In: Berg JC, ed. Wettability. New York: Marcel Dekker; 1993:2–74.
19.Petrucci RH, Harwood WS, Herring FG. General Chemistry: Principles and Modern Applications. Upper Saddle River, NJ: Prentice Hall; 2002.
20.Fatt I. Prentice Medal lecture: contact lens wettability—myths, mysteries, and realities. Am J Optom Physiol Opt 1984;61:419–30.
21.Port MJ. Contact lens surface properties and interactions. Optom Today 1999;39:27–35.
22.Bruce AS, Mainstone JC, Golding TR. Analysis of tear film breakup on etafilcon A hydrogel lenses. Biomaterials 2001;22:3249–56.
23.Carney F, Keay L, Stapleton F, Morris C, Willcox M. Hydrogel lens wettability and deposition in vivo. Clin Exp Optom 1998;81:2:51–5.
24.Doane MG. An instrument for in vivo tear film interferometry. Optom Vis Sci 1989;66:383–8.
25.Guillon M, Guillon JP. Hydrogel lens wettability during overnight wear. Ophthal Physiol Opt 1989;9:355–9.
26.Nichols JJ, King-Smith PE. Thickness of the pre- and post-contact lens tear film measured in vivo by interferometry. Invest Ophthalmol Vis Sci 2003;44:68–77.
27.Nichols JJ, Mitchell GL, King-Smith PE. Thinning rate of the precorneal and prelens tear films. Invest Ophthalmol Vis Sci 2005;46:2353–61.
28.Cheng L, Muller SJ, Radke CJ. Wettability of silicone-hydrogel contact lenses in the presence of tear-film components. Curr Eye Res 2004;28:93–108.
29.Pethica BA. The physical chemistry of cell adhesion. Exp Cell Res 1961;suppl 8:123–40.
30.Holly FJ, Refojo MF. Wettability of hydrogels. I. Poly (2-hydroxyethyl methacrylate). J Biomed Mater Res 1975;9:315–26.
31.Samoilova NA, Krayukhina MA, Novikova SP, Babushkina TA, Volkov IO, Komarova LI, Moukhametova LI, Aisina RB, Obraztsova EA, Yaminsky IV, Yamskov IA. Polyelectrolyte thromboresistant affinity coatings for modification of devices contacting blood. J Biomed Mater Res A 2007;82:589–98.
32.Hoque E, DeRose JA, Hoffmann P, Bhushan B, Mathieu HJ. Chemical stability of nonwetting, low adhesion self-assembled monolayer films formed by perfluoroalkylsilanization of copper. J Chem Phys 2007;126:114706.
33.Maldonado-Codina C, Morgan PB. In vitro water wettability of silicone hydrogel contact lenses determined using the sessile drop and captive bubble techniques. J Biomed Mater Res A 2007;83:496–502.
34.Taylor M, Urquhart AJ, Zelzer M, Davies MC, Alexander MR. Picoliter water contact angle measurement on polymers. Langmuir 2007;23:6875–8.
35.Lorentz H, Rogers R, Jones L. The impact of lipid on contact angle wettability. Optom Vis Sci 2007;84:946–53.
36.Lleixa Calvet J, Grafahrend D, Klee D, Moller M. Sterilization effects on starPEG coated polymer surfaces: characterization and cell viability. J Mater Sci Mater Med 2008;19:1631–6.
37.Aguilar-Mendoza JA, Rosales-Leal JI, Rodriguez-Valverde MA, Gonzalez-Lopez S, Cabrerizo-Vilchez MA. Wettability and bonding of self-etching dental adhesives. Influence of the smear layer. Dent Mater 2008;24:994–1000.
38.Goswami S, Klaus S, Benziger J. Wetting and absorption of water drops on Nafion films. Langmuir 2008;24:8627–33.
39.Drelich J, Miller JD, Good RJ. The effect of drop (bubble) size on advancing and receding contact angles for heterogeneous and rough solid surfaces as observed with sessile-drop and captive-bubble techniques. J Colloid Interface Sci 1996;179:37–50.
40.Zhang W, Hallstrom B. Membrane characterization using the contact angle technique I. Methodology of the captive bubble technique. Desalination 1990;79:1–12.
41.Krishnan A, Liu YH, Cha P, Woodward R, Allara D, Vogler EA. An evaluation of methods for contact angle measurement. Colloids Surf B Biointerfaces 2005;43:95–8.
42.Tonge S, Jones L, Goodall S, Tighe B. The ex vivo wettability of soft contact lenses. Curr Eye Res 2001;23:51–9.
43.Read M, Maldonado-Codina C, Morgan P. The repeatability of contact angle measurements on hydrogel materials. CLAO J 2008;31:253–4.
44.Shang J, Flury M, Harsh JB, Zollars RL. Comparison of different methods to measure contact angles of soil colloids. J Colloid Interface Sci 2008;328:299–307.
45.Han DK, Park KD, Ryu GH, Kim UY, Min BG, Kim YH. Plasma protein adsorption to sulfonated poly(ethylene oxide)-grafted polyurethane surface. J Biomed Mater Res 1996;30:23–30.
46.Desai NP, Hubbell JA. Biological responses to polyethylene oxide modified polyethylene terephthalate surfaces. J Biomed Mater Res 1991;25:829–43.
47.Tzoneva R, Faucheux N, Groth T. Wettability of substrata controls cell-substrate and cell-cell adhesions. Biochim Biophys Acta 2007;1770:1538–47.
48.Kim MS, Shin YN, Cho MH, Kim SH, Kim SK, Cho YH, Khang G, Lee IW, Lee HB. Adhesion behavior of human bone marrow stromal cells on differentially wettable polymer surfaces. Tissue Eng 2007;13:2095–103.
49.Norman JJ, Desai TA. Control of cellular organization in three dimensions using a microfabricated polydimethylsiloxane-collagen composite tissue scaffold. Tissue Eng 2005;11:378–86.
50.Nishio R, Nakayama M, Ikekita M, Watanabe Y. Auxiliary liver organ formation by implantation of spleen-encapsulated hepatocytes. Tissue Eng 2006;12:2565–72.
51.Gumbiner BM. Cell adhesion: the molecular basis of tissue architecture and morphogenesis. Cell 1996;84:345–57.
52.Harnett EM, Alderman J, Wood T. The surface energy of various biomaterials coated with adhesion molecules used in cell culture. Colloids Surf B Biointerfaces 2007;55:1:90–7.
53.Underwood PA, Bean PA, Gamble JR. Rate of endothelial expansion is controlled by cell:cell adhesion. Int J Biochem Cell Biol 2002;34:55–69.
54.Faucheux N, Schweiss R, Lutzow K, Werner C, Groth T. Self-assembled monolayers with different terminating groups as model substrates for cell adhesion studies. Biomaterials 2004;25:2721–30.
55.Arima Y, Iwata H. Effect of wettability and surface functional groups on protein adsorption and cell adhesion using well-defined mixed self-assembled monolayers. Biomaterials 2007;28:3074–82.
56.Courtney JM, Lamba NM, Sundaram S, Forbes CD. Biomaterials for blood-contacting applications. Biomaterials 1994;15:737–44.
57.Tzoneva R, Groth T, Altankov G, Paul D. Remodeling of fibrinogen by endothelial cells in dependence on fibronectin matrix assembly. Effect of substratum wettability. J Mater Sci Mater Med 2002;13:1235–44.
58.Wan GJ, Huang N, Yang P, Fu RK, Ho JP, Xie X, Zhou HF, Chu PK. Platelet activation behavior on nitrogen plasma-implanted silicon. Mater Sci Eng C 2007;27:928–32.
59.Yang P, Huang N, Leng YX, Yao ZQ, Zhou HF, Maitz M, Leng Y, Chu PK. Wettability and biocompatibility of nitrogen-doped hydrogenated amouphous carbon films: effect of nitrogen. Nuclear Instruments Meth Phys Res B 2006;242:22–25.
60.Deppisch R, Storr M, Buck R, Gohl H. Blood material interactions at the surfaces of membranes in medical applications. Sep Purif Tech 1998;14:241–54.
61.Okano T, Aoyagi T, Kataoka K, Abe K, Sakurai Y, Shimada M, Shinohara I. Hydrophilic-hydrophobic microdomain surfaces having an ability to suppress platelet aggregation and their in vitro antithrombogenicity. J Biomed Mater Res 1986;20:919–27.
62.Halverson GE, Nelson GA. Dental impression supply kit. US patent 4,763,791. August 16, 1988.
63.Michalakis KX, Bakopoulou A, Hirayama H, Garefis DP, Garefis PD. Pre- and post-set hydrophilicity of elastomeric impression materials. J Prosthodont 2007;16:238–48.
64.Kess RS, Combe EC, Sparks BS. Effect of surface treatments on the wettability of vinyl polysiloxane impression materials. J Prosthet Dent 2000;84:98–102.
65.Abdelaziz KM, Combe EC, Hodges JS. The wetting of surface-treated silicone impression materials by gypsum mixes containing disinfectants and modifiers. J Prosthodont 2005;14:104–9.
66.Lloyd AW, Faragher RG, Denyer SP. Ocular biomaterials and implants. Biomaterials 2001;22:769–85.
67.Pearce JL. Intraocular lenses. Curr Opin Ophthalmol 1990;1:42–8.
68.Tognetto D, Toto L, Minutola D, Ballone E, Di Nicola M, Di Mascio R, Ravalico G. Hydrophobic acrylic versus heparin surface-modified polymethylmethacrylate intraocular lens: a biocompatibility study. Graefes Arch Clin Exp Ophthalmol 2003;241:625–30.
69.Rosen E. Intraocular lenses. Curr Opin Ophthalmol 1994;5:40–54.
70.Hollick EJ, Spalton DJ, Ursell PG, Pande MV. Biocompatibility of poly(methyl methacrylate), silicone, and AcrySof intraocular lenses: randomized comparison of the cellular reaction on the anterior lens surface. J Cataract Refract Surg 1998;24:361–6.
71.Pande MV, Spalton DJ, Kerr-Muir MG, Marshall J. Cellular reaction on the anterior surface of poly(methyl methacrylate) intraocular lenses. J Cataract Refract Surg 1996;22(suppl 1):811–7.
72.Ohnishi Y, Yoshitomi T, Sakamoto T, Fujisawa K, Ishibashi T. Evaluation of cellular adhesions on silicone and poly(methyl methacrylate) intraocular lenses in monkey eyes: an electron microscopic study. J Cataract Refract Surg 2001;27:2036–40.
73.Amon M, Menapace R, Radax U, Freyler H. In vivo study of cell reactions on poly(methyl methacrylate) intraocular lenses with different surface properties. J Cataract Refract Surg 1996;22(suppl 1):825–9.
74.Amon M, Menapace R. Long-term results and biocompatibility of heparin-surface-modified intraocular lenses. J Cataract Refract Surg 1993;19:258–62.
75.Mester U, Strauss M, Grewing R. Biocompatibility and blood-aqueous barrier impairment in at-risk eyes with heparin-surface-modified or unmodified lenses. J Cataract Refract Surg 1998;24:380–4.
76.Pearce JL. Intraocular lenses. Curr Opin Ophthalmol 1992;3:29–38.
77.Yao K, Huang XD, Huang XJ, Xu ZK. Improvement of the surface biocompatibility of silicone intraocular lens by the plasma-induced tethering of phospholipid moieties. J Biomed Mater Res A 2006;78:684–92.
78.Schroeder AC, Lingenfelder C, Seitz B, Grabowy U, W Spraul C, Gatzioufas Z, Herrmann M. Impact of fibronectin on surface properties of intraocular lenses. Graefes Arch Clin Exp Ophthalmol 2009;247:1277–83.
79.Lever OW, Groemminger SF, Allen ME, Bornemann RH, Dey DR, Barna BJ. Evaluation of the relationship between total lens protein deposition and patient-rated comfort of hydrophilic (soft) contact lenses. Int Contact Lens Clin 1995;22:5–13.
80.Guryca V, Hobzova R, Pradny M, Sirc J, Michalek J. Surface morphology of contact lenses probed with microscopy techniques. Cont Lens Anterior Eye 2007;30:215–22.
81.Dumbleton KA, Chalmers RL, Richter DB, Fonn D. Vascular response to extended wear of hydrogel lenses with high and low oxygen permeability. Optom Vis Sci 2001;78:147–51.
82.Tighe B. Silicone hydrogel materials—how do they work? In: Sweeney DF, ed. Silicone Hydrogels—The Rebirth of Continuous Wear Contact Lenses. Oxford, UK: Butterworth-Heinemann; 2000:1–21.
83.Tighe B. Silicone hydrogels: structure, properties and behaviour: In: Sweeney D, ed. Silicone Hydrogels: Continuous Wear Contact Lenses, 2nd ed. Oxford, UK: Butterworth-Heinemann; 2004:1–27.
84.Nicolson PC. Continuous wear contact lens surface chemistry and wearability. Eye Contact Lens 2003;29:S30–2.
85.Lopez-Alemany A, Compan V, Refojo MF. Porous structure of Purevision versus Focus Night & Day and conventional hydrogel contact lenses. J Biomed Mater Res 2002;63:319–25.
86.Gonzalez-Meijome JM, Lopez-Alemany A, Almeida JB, Parafita MA, Refojo MF. Microscopic observation of unworn siloxane-hydrogel soft contact lenses by atomic force microscopy. J Biomed Mater Res B Appl Biomater 2006;76:412–8.
87.Kunzler J. Silicone-based hydrogels for contact lens applications. Contact Lens Spectrum 1999;14(suppl 8):9–11.
88.Nicolson PC, Baron RC, Chabrecek P, Court JL, Domschke A, Griesser HJ, Ho A, Hopken J, Laycock B, Liu Q, Lohmann D, Meijs GF, Papaspiliotopoulos E, Riffle JS, Schindhelm K, Sweeney D, Terry WL Jr, Vogt J, Winterton LC. Extended wear ophthalmic lens. US patent 5,760,100. June 2,1998.
89.Teichroeb JH, Forrest JA, Ngai V, Martin JW, Jones L, Medley J. Imaging protein deposits on contact lens materials. Optom Vis Sci 2008;85:1151–64.
90.Steffen R, K McCabe. Finding the comfort zone. Contact Lens Spectrum 2004;13(suppl 3):1–4.
91.Jones L, Subbaraman LN, Rogers R, Dumbleton KA. Surface treatment, wetting and modulus of silicone hydrogels. Optician 2006;232:28–33.
92.Jones L. A new silicone hydrogel lens comes to market. Contact Lens Spectrum 2007;22:23–4.
93.Jones L. Comfilcon A: a new silicone hydrogel material. Contact Lens Spectrum 2007;22:21.
94.Cheung SW, Cho P, Chan B, Choy C, Ng V. A comparative study of biweekly disposable contact lenses: silicone hydrogel versus hydrogel. Clin Exp Optom 2007;90:124–31.
95.Keir N, Boone A, Jones L, Woods CA, Fonn D. In vivo and ex vivo wettability and the association with contact lens comfort. Cont Lens Anterior Eye 2008;31:6:292.
96.Dumbleton KA, Woods CA, Jones LW, Fonn D. Comfort and adaptation to silicone hydrogel lenses for daily wear. Eye Contact Lens 2008;34:215–23.
97.Morgan PB, Efron N. Comparative clinical performance of two silicone hydrogel contact lenses for continuous wear. Clin Exp Optom 2002;85:183–92.
98.Tripathi RC, Tripathi BJ, Ruben M. The pathology of soft contact lens spoilage. Ophthalmology 1980;87:365–80.
99.Gellatly KW, Brennan NA, Efron N. Visual decrement with deposit accumulation of HEMA contact lenses. Am J Optom Physiol Opt 1988;65:937–41.
100.Pritchard N, Fonn D, Weed K. Ocular and subjective responses to frequent replacement of daily wear soft contact lenses. CLAO J 1996;22:53–9.
101.Jones L, Senchyna M, Glasier MA, Schickler J, Forbes I, Louie D, May C. Lysozyme and lipid deposition on silicone hydrogel contact lens materials. Eye Contact Lens 2003;29:S75–9.
102.Santos L, Rodrigues D, Lira M, Oliveira ME, Oliveira R, Vilar EY, Azeredo J. The influence of surface treatment on hydrophobicity, protein adsorption and microbial colonisation of silicone hydrogel contact lenses. Cont Lens Anterior Eye 2007;30:183–8.
103.Garrett Q, Laycock B, Garrett RW. Hydrogel lens monomer constituents modulate protein sorption. Invest Ophthalmol Vis Sci 2000;41:1687–95.
104.Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, Lappin-Scott HM. Microbial biofilms. Annu Rev Microbiol 1995;49:711–45.
105.Costerton JW, Cheng KJ, Geesey GG, Ladd TI, Nickel JC, Dasgupta M, Marrie TJ. Bacterial biofilms in nature and disease. Annu Rev Microbiol 1987;41:435–64.
106.Mercuri LG. Microbial biofilms: a potential source for alloplastic device failure. J Oral Maxillofac Surg 2006;64:1303–9.
107.Tapia G, Yee J. Biofilm: its relevance in kidney disease. Adv Chronic Kidney Dis 2006;13:215–24.
108.Anguita-Alonso P, Hanssen AD, Patel R. Prosthetic joint infection. Expert Rev Anti Infect Ther 2005;3:797–804.
109.Hall-Stoodley L, Costerton JW, Stoodley P. Bacterial biofilms: from the natural environment to infectious diseases. Nat Rev Microbiol 2004;2:95–108.
110.Tang H, Cao T, Liang X, Wang A, Salley SO, McAllister J II, Ng KY. Influence of silicone surface roughness and hydrophobicity on adhesion and colonization of Staphylococcus epidermidis. J Biomed Mater Res A 2009;88:454–63.
111.Quirynen M, van der Mei HC, Bollen CM, Schotte A, Marechal M, Doornbusch GI, Naert I, Busscher HJ, van Steenberghe D. An in vivo study of the influence of the surface roughness of implants on the microbiology of supra- and subgingival plaque. J Dent Res 1993;72:1304–9.
112.Higashi JM, Wang IW, Shlaes DM, Anderson JM, Marchant RE. Adhesion of Staphylococcus epidermidis and transposon mutant strains to hydrophobic polyethylene. J Biomed Mater Res 1998;39:341–50.
113.Okada A, Nikaido T, Ikeda M, Okada K, Yamauchi J, Foxton RM, Sawada H, Tagami J, Matin K. Inhibition of biofilm formation using newly developed coating materials with self-cleaning properties. Dent Mater J 2008;27:565–72.
114.Henriques M, Sousa C, Lira M, Elisabete M, Oliveira R, Azeredo J. Adhesion of Pseudomonas aeruginosa and Staphylococcus epidermidis to silicone-hydrogel contact lenses. Optom Vis Sci 2005;82:446–50.
115.Kodjikian L, Burillon C, Roques C, Pellon G, Freney J, Renaud FN. Bacterial adherence of Staphylococcus epidermidis to intraocular lenses: a bioluminescence and scanning electron microscopy study. Invest Ophthalmol Vis Sci 2003;44:4388–94.
116.Bruinsma GM, van der Mei HC, Busscher HJ. Bacterial adhesion to surface hydrophilic and hydrophobic contact lenses. Biomaterials 2001;22:3217–24.
117.Vermeltfoort PB, van der Mei HC, Busscher HJ, Hooymans JM, Bruinsma GM. Physicochemical factors influencing bacterial transfer from contact lenses to surfaces with different roughness and wettability. J Biomed Mater Res B Appl Biomater 2004;71:336–42.
This article has been cited 4 time(s).
LangmuirA Robust Polynomial Fitting Approach for Contact Angle MeasurementsLangmuir
Plos OneRisk Factors for Acute Endophthalmitis following Cataract Surgery: A Systematic Review and Meta-AnalysisPlos One
Journal of Biomedical Materials Research Part ABiofunctionalization of titanium surface with multilayer films modified by heparin-VEGF-fibronectin complex to improve endothelial cell proliferation and blood compatibilityJournal of Biomedical Materials Research Part A
Rsc AdvancesElectrophoretic deposition of nanostructured-TiO2/chitosan composite coatings on stainless steelRsc Advances
biomaterials; biocompatibility; wettability; contact angle; hydrophilic; hydrophobic
© 2010 American Academy of Optometry
Highlight selected keywords in the article text.