Towards proteomics-on-chip: The role of the surface

Abstract

Miniaturisation is revolutionary to high-throughput proteomics. These technologies have gained much interest in the past decade, as they allow for sensitive parallel analysis of small amounts of biological materials. This review describes the state of the art of proteomics-on-chip, with a particular focus on the fundamental proteomics-on-chip challenges. The important role of bio-interfacial interactions and strategies to control them are presented. Various coating methodologies for on-chip protein/peptide separation are reviewed to provide an overview of the principles of protein-resistant and protein immobilisation coatings, and their effectiveness.

---

Malinda Salim

Malinda Salim received her BEng degree in Chemical Engineering from the University of Sheffield in 2004. She pursued her PhD studies in developing surface modification techniques for proteomics-on-chip application with joined collaboration between Chemical Engineering Department and Engineering Materials, under the supervision of Professor Phillip Wright and Dr Sally McArthur. Upon completing her PhD, she took up a postdoctoral research associate position with Professor Phillip Wright and Dr Seetharaman (Raman) Vaidyanathan at the University of Sheffield, applying her multi-disciplinary research background to metabolomics. She is particularly interested in the application of mass spectrometries to investigate metabolites interaction.

Sally L. McArthur

Sally McArthur is currently Associate Professor in Bioengineering in the Faculty of Engineering and Industrial Science at Swinburne University of Technology in Australia. Prior to that she in the Department of Engineering Materials at the University of Sheffield as Senior Lecturer in Biomedical Engineering after joining the department as a Lecturer in May 2002. Prior to moving to the UK, she was a Senior Research Fellow in the Bioengineering Department at the University of Washington, working with Professor David Castner and NESAC/Bio.

S. Vaidyanathan

Raman Vaidyanathan joined the Department Chemical and Process Engineering at The University of Sheffield in 2007, as a lecturer. He's a chemical graduate with a Masters degree in Biotechnology, both from India. He completed his PhD in 2001 from the University of Strathclyde in Glasgow, where he investigated the application of near infrared spectroscopy to monitor industrially relevant bioprocesses involving filamentous microorganisms. This was work done in collaboration with Eli Lilly. He has over five years of pre-doctoral industrial experience applying biotechnological solutions to environmental engineering problems.

Phillip C. Wright

Phillip Wright is the foundation Professor of Systems Biology and Engineering within the Department of Chemical and Process Engineering (CPE), at The University of Sheffield. He is also the Head of Department, ad Director of the ChELSI Institute at Sheffield. He started his Chemical Engineering career as a cadet at BHP Steel in Wollongong NSW Australia, and completed his BE in Chemical Engineering by a combination of both full and part time study, graduating from the University of NSW in 1991. He returned to BHP Steel as a Process Engineer for the years 1991–1993.

---

1. Introduction to miniaturisation and proteomics

1.1 Miniaturisation

The first study of miniaturisation was performed in 1975 by Terry et al. on a microfabricated silicon gas chromatographic analyser.¹ However, research and development of lab-on-a-chip applications boomed in the 1990's following the report by Manz et al. who proposed miniaturised devices for chemical analysis.² Miniaturisation has demonstrated advantages over macroscale systems due to lower sample consumption, rapid high throughput analysis, and possibilities for integration and automation (including scale-out). Reduced sample consumption is desirable to save precious samples and chemicals. Miniaturisation can also shorten the time to obtain the result, since this architecture has the possibility to run multiple samples simultaneously via parallelisation, dramatically increasing throughput.^3–5 Although the nature of the reaction does not usually change in miniaturised devices, the diffusion time (t) becomes significantly shorter,^3,6 since t is proportional to the square of the characteristic length, d (i.e. t α d²). An immunoassay performed in the usual 96-well plate requires hours for an antibody-antigen reaction to reach optimal levels, and only minutes or seconds in a miniaturised immunoassay.^5,7,8 Another change upon scaling down is the higher surface area-to-volume ratio, which accounts for the important role of surface characteristics, hence the control of biomolecules and electroosmotic flow (EOF) in miniaturised systems.^4,5,9

The advantages of miniaturisation, together with an ability to integrate several experimental steps into a single micro-scale device, make microfluidics a promising tool for biological analyses.^4,5,10 Recently, microfluidic devices were demonstrated as being able to mimic biological systems within the body, enabling in vivo studies to be undertaken in an in vitro environment. Key applications have included the imitation of blood vessels,¹¹ kidney,¹² and lung.¹³Cell culture has also been performed in microchannels to better understand effects of shear and diffusion on cell behavior in vivo.¹⁴

1.2 Proteomics

Proteomics (proteome–protein complement expressed by the genome)¹⁵ is important for pharmaceutical, biomedical, environmental and biotechnology applications.^16,17 Core proteomic analytical processes usually include protein extraction from cells, protein separation, protein digestion (with a protease: usually with trypsin into peptides in what is known as “bottom up” proteomics), and analysis using mass spectrometry (MS).^16,18–20 As these conventional processes can be slow, labour intensive, and introduce sample loss and degradation in each processing stage, miniaturisation and integration of these processes onto a microfluidic chip, usually coupled to a mass spectrometer are being widely investigated to improve throughput and detection.^16,18–20 Two-dimensional sodium dodecyl sulfate poly(acrylamide) gel electrophoresis (2D SDS-PAGE) is a common technology used in proteomics for orthogonal protein separation based on isoelectric points and molecular weights. It usually requires several hours (often overnight) for completion,²¹ which reduces to minutes in miniaturised 2D SDS-PAGE apparatus.^22,23

To develop a realistic proteomics-on-chip device, a diverse set of research fields need to be brought together. These can be divided into categories such as micromachining and microfluidic processes; protein–surface interactions; surface modification; proteomics workflows (protein extraction from cells, protein digestion, protein and peptide separations); coupling between unit operations; and detection strategies. Each category has its own set of intertwined challenges. For example, effective adaptations of commonly used cell lysis techniques (e.g. mechanically-based processes such as grinding in liquid nitrogen with a mortar and pestle), are difficult to achieve on chip.²⁴ Others, such as the efficacy of proteolytic digestion on-chip, depend heavily on the success of enzyme-surface immobilisation (often reported to provide higher efficiency compared to conventional ‘in-solution’ digestion),²⁵ preserving enzymatic activity, sample solubility, and product recovery. Packing of solid materials including beads and gels in microchannels has also been shown to be challenging.²⁶ All of the issues require dedicated research.

Downstream, on chip devices must also ensure compatibility with (mostly electrospray ionisation (ESI) based) MS post-analysis. Readers interested in applications and methods to interface microfluidic chip-MS (ESI and MALDI) are directed to recent reviews by Jeonghoon et al.^27,28 General concerns include the influence of current from the separation chamber on the electrospray current, and chip-interface design.^29,30 High EOF, the bulk flow of fluid through the channel brought about by an electric field, is usually also required (e.g. using a sheathless interface).³¹ High EOF is, however, undesirable for many forms of protein separation.^30,32 Additionally, band broadening resulting from Joule heating can detrimentally affect the separation resolution.³³ Biomolecule adsorption to channel walls has also been shown to significantly affect the EOF,^34,35 but many of the typically polyethylene oxide (PEO)-based low fouling coatings exhibit low EOF.^35,36

Surfaces, biomolecules, and fluid movement are therefore ubiquitous and inter-dependent within any on-chip device. Controlling the extent of EOF while simultaneously controlling biointerfacial interactions through surface modification is thus critical to the development of effective on-chip proteomic devices.^19,37 Different surface properties can provide different controls for EOF, prevent non-specific adsorption of biomolecules, or enable the immobilisation of biomolecules for specific capture, digestion or detection methods. Given the multidisciplinary nature of this field, there are many other issues related to on-chip proteomics that are beyond the scope of this review. A number of articles however, are recommended to readers who require more information.^{19,31,38–42}

1.3 Scope

This review seeks to integrate fundamental proteomics-on-chip challenges to provide a readily accessible overview. The first part (section 2) of this article presents a primer on biomolecule-surface interactions, focusing on the mechanisms and impact of uncontrolled adsorption in microfluidic devices. This provides the knowledge required to understand measures that can be taken to control biointerfacial interactions within microfluidic devices for biomolecule separation using a range of surface modification strategies in section 3. Perspectives are given on the current technologies and future prospects. Finally, in section 4, a critical evaluation of surfaces that resist non-specific bioadhesion, in particular PEO-based surfaces, is included to provide an overview of the principles of protein-resistant coatings and the coating effectiveness of various common strategies.

2. Understanding protein adsorption at solid–liquid interface

2.1 Interaction between proteins and surfaces

Proteins are complex heterogeneous biological molecules that are composed of chains of amino acids (NH₂–CHR–COOH) which can be hydrophobic or hydrophilic, neutral, negatively or positively charged.⁴³ As a consequence, proteins are able to interact with each other as well as synthetic surfaces in a number of ways as shown in Fig. 1. These include hydrophobic interactions, repulsive and attractive ionic or electrostatic interactions, and short range interactions such as hydrogen bonding and van der Waals interactions.^43,44 A delicate balance between a number of forces plays an important role in the stabilisation of a protein, which may be perturbed upon adsorption to or interactions with a solid surface. Since protein adsorption normally occurs at constant temperature and pressure, the thermodynamic driving forces governing them can be described by considering the Gibbs free energy for the system:

ΔG_ads = ΔH_ads − TΔS_ads

where G, H, S and T are Gibbs free energy, enthalpy, entropy and absolute temperature respectively. Generally, if the Gibbs free energy of the system is negative (ΔG_ads < 0), proteins will adsorb onto a surface. For a negative ΔG_ads, there should either be a decrease in enthalpy or an increase in entropy.^43,45 Norde reported that the entropy gained from the protein unfolding upon adsorption onto surfaces may be the major driving force for protein adsorption.^43,45

Fig. 1 Schematic representation of protein interaction (having hydrophobic, neutral hydrophilic and charged sides) with a number of synthetic surfaces [Adapted from ref. 46].

2.2.1 Hydrophobic interactions. Hydrophobic interaction is an additional driving force for protein adsorption at surfaces. This interaction may be short range (ca. 15–20 nm), although the magnitude of hydrophobic interaction (long or short range) is still under debate. Hydrophobic interaction minimises the free energy of the system by reducing the interfacial area between non-polar regions and aqueous solution.⁴⁷ This gives rise to protein adsorption when the protein is brought into contact with a hydrophobic surface. The hydrophobic moieties residing in the protein interior may therefore be exposed to the hydrophobic surface, which cause protein unfolding. Therefore, protein adsorption onto a hydrophobic surface generally results in changes of the protein structure or denaturation (Fig. 2). The extent of denaturation depends on the degree of hydrophobicity of the protein and the surface.^43,44,48,49

Fig. 2 Schematic representation of protein unfolding on a hydrophobic surface.

Several groups have reported the effects of surface hydrophobicity on protein denaturation. For example, Wertz et al. studied the spreading of bovine serum albumin (BSA) and human fibrinogen on hydrophobic and hydrophilic self-assembled monolayers (SAMs) using total internal reflection fluorescence.⁵⁰ For both proteins, fast and extensive spreading was observed on the hydrophobic surface (greater increase of protein footprint as a function of time), whereas very little spreading was observed on the hydrophilic surface. A protein footprint is the surface area a protein occupies when adsorbed onto a surface. The ultimate surface coverage for both proteins was found to be smaller on hydrophobic compared to hydrophilic surfaces at high protein concentrations. Similarly, as reported by Veen et al., the adsorbed amount of lysozyme and α-lactalbumin at their saturation level is smaller on hydrophobic surfaces compared to hydrophilic surfaces at high protein concentrations.⁵¹ At low protein concentrations, however, the quantity of adsorbed proteins was greater on a hydrophobic surface. Although faster spreading of protein occurred on the hydrophobic surface, the proteins spread to a far lesser extent at low protein concentration. These studies contradict several general findings that greater protein surface coverage was obtained on hydrophobic surfaces, where protein spreading is usually not accounted for.^52–55 The aforementioned studies also showed the dependency of protein spreading on protein supply rate; and that a balance exists between the rate of protein spreading and the rate of protein adsorption.^50,51,56

2.2.2 Ionic interactions. Ionic/electrostatic interactions arise from the attractive and repulsive interactions between the charge groups on the proteins, and those present on the surface.⁴⁴ The net charges of the protein and surface are affected by the pH and the ionic strength of the solution, defined by the Debye length (between 1–1000 nm).⁵⁷ These short and long range electrostatic interactions also play an important role in protein stability and function.⁵⁸ A charged protein is expected to be attracted or adsorbed onto an oppositely charged surface, and be repelled when both the protein and the surface are similarly charged, if electrostatic interactions dominate. This, however, is not always the case, as adsorption of proteins occurs even on similarly charged surfaces. For example, net negatively charged proteins have been shown to adsorb onto negatively charged surfaces.^43,45,48,59 Although a protein carries a net charge, it cannot be considered as a single point charge, as different amino acid residues exist on different parts of the protein surface,⁶⁰ creating a charge distribution across the surface.

2.2.3 Other interactions. The role of van der Waals forces in protein–surface interactions is of importance when the protein is in close contact with the surface, as van der Waals interactions are short ranged (typically below 2.5 nm) and always attractive.⁶¹ This interaction can be formed between polar or non-polar atoms or molecules. The balance between van der Waals and electrostatic interactions for colloidal or protein systems and stability has been interpreted in the Derjaguin-Landau-Verwey-Overbeek (DLVO) theory.⁶²

Hydrogen bonding, which also has a short range character (ca. 0.3 nm), is another type of interaction between a protein and a solid surface.⁶³ It is formed when a proton is shared between two electronegative atoms. Therefore, H-bonding may form between amino acids (intramolecular hydrogen bonding, which also contributes to protein stability) or between a protein and surrounding water molecules,^63,64 which can stabilise native protein conformation and preserve its activity.

2.2 Factors that influence protein adsorption

Understanding of the principles and factors that influence protein adsorption is important for several applications including biomaterials, pharmaceuticals, food technology, biotechnologies, filtration and biosensors.⁴⁴Protein adsorption at surfaces is a complex phenomenon, and is dependent on factors such as protein properties, protein orientation on the surface, the surrounding environment, and surface properties.^{43–45,48,65,66} Examples of the properties of the proteins that affect adsorption are the charge, size, and structural stability. The extent of the interaction between a protein and the surface will generally be greater if the protein is large, carries zero net charge, and/or has low structural stability. A large protein, for example, human fibrinogen (340 kDa, 5 nm × 5 nm × 47 nm),⁵⁴ has the ability to form multiple contact points with the surface, while smaller proteins such as chicken-egg lysozyme (14.5 kDa, 4.5 nm × 3 nm × 3 nm)⁶⁷ covers a smaller area, and thus can have fewer contact points per molecule.^{44,45,48,49,62,68} When the protein has zero net charge (i.e. near its isoelectric point, pI), the reduced electrostatic repulsion between the individual proteins will allow more molecules to be adsorbed onto a surface. Finally, a protein that has low structural stability can readily undergo denaturation or ‘spreading’. This allows more sites on the protein to be in contact with the surface, creating greater surface interaction (see Fig. 2).^{43,44,48,67,68}

The properties of surfaces such as surface topography, composition, charge, and heterogeneity will also determine the protein–surface interaction. A surface having rough topography will have a greater surface area available for the attachment of the protein. On surfaces with heterogeneous chemical compositions, different types of protein interaction may occur, depending on the density and distribution of functional and non-functional groups of the protein and the surface. In short, increasing the surface complexities may increase the types and numbers of interactions between the protein and the surface. Other than all these properties, the surrounding environment factors such as protein concentrations, time, pH of the media, flow conditions, ionic strengths of the buffer and temperature will also affect the protein adsorption behaviour.^43,44,49,69

2.3 Methods of measuring protein adsorption

This section describes some of the techniques that have been developed to investigate protein adsorption at surfaces: both planar and microchannels. These can generally be divided into two classes, i.e. surface analytical and (bio) chemical (see Table 1). Although a wide range of surface characterisation methods exists, measurement of protein adsorption in closed-system microchannels is observed to be often limited to (bio)chemical analysis techniques such as radiochemical (most commonly isotopic labelling), immunological, fluorescence, and electrophoretic due to the difficulty in instrumental miniaturisation and assessing the inner surface of the microchannel. Monitoring of protein adsorption has however been shown possible in microfluidic chips integrated to surface plasmon resonance and surface plasmon resonance imaging.^70,71 The information obtained and limitations of each of these techniques are presented in Table 1, where in all cases, these approaches provide minimal data on the structure of adsorbed protein layers.

Table 1 General techniques used to characterise protein adsorption^53,72–83

Method	Information	Limitations
Surface analytical
Atomic force microscopy	Interaction forces, structure, kinetics	Protein distortion due to sample-tip interaction, difficult sample preparation, tip contamination, not quantitative.
Circular dichroism	Structure	Signal noise problems from interfering compounds.
Contact angle measurement	Wetting, hydrophilicity/hydrophobicity	Not quantitative, limited adsorption information.
Ellipsometry	Quantitative, structure, kinetics	Surface refractive index difficult to determine.
Reflectometry	Quantitative, structure, kinetics	Surface refractive index difficult to determine.
Secondary ion mass spectrometry with time-of-flight	Structure, chemical	Dehydrated state, vacuum, complex data, difficulty in identifying individual proteins, largest protein fragments are the individual amino acid immonium ions.
Surface plasmon resonance	Quantitative, kinetics	Analytes must have sufficient mass (>2000 Daltons).
Surface-matrix assisted laser desorption/ionisation mass spectrometry	Semi quantitative, molecular mass of adsorbed protein	No information on the structure of adsorbed protein layer, dehydrated state, vacuum, incorporation of protein into matrix difficult.
Total internal reflection fluorescence	Quantitative, structure, kinetics	Requires fluorescent labeling.
X-ray photoelectron spectroscopy	Quantitative	Little detail on molecular information, identify different proteins difficult, vacuum, dehydrated state. May cause denaturation and alter protein film thickness.
Biochemical
Enzyme linked immunosorbent assay	Kinetics	Not quantitative. Protein orientation/conformation and antibody dependent. Limited types of proteins depending on the antibodies available.
Fluorescence spectroscopy	Quantitative, kinetics	No information on the structure of adsorbed protein layer, labeling affects protein.
SDS-PAGE	Quantitative	Low sensitivity (dependent on intensity of protein bands/spots), poor reproducibility.
Electrophoresis	Semi quantitative, kinetics	No information on the structure of adsorbed protein layer, not quantitative.
Radiochemical	Quantitative, kinetics	No information on the structure of adsorbed protein layer, environmental issue on disposal, labeling affects protein.

---

3. Protein adsorption and surface modification strategies in microchannel

3.1 Protein adsorption

There are thousands of peer reviewed articles relating to protein adsorption onto non-microchannel surfaces, which incorporate many adsorption variations such as different types of proteins, surfaces chemistries and structures, protein adsorption properties, and characterisation methods.^84–90 Since the realisation of the importance of miniaturisation in biotechnology, reports on protein adsorption behaviour in microchannel systems have also emerged.^8,83,91–95 Many publications however focus on system design and applications such as varying surface treatments for controlling microchannel properties, which will be discussed further in section 3.2.

Rossier et al. studied the adsorption isotherm of a protein and its antibody, as well as the immunosorption kinetics onto polymeric microchannels using a radiolabelling method.⁸ They observed fast diffusion of molecules onto the channel wall, with complete immunoreaction taking place in 1 min. However, the radiolabelling process is not desirable for studying protein adsorption as it is expensive and hazardous.⁹⁶ In 2003, Lenghaus et al. reported the first attempt to elucidate enzyme adsorption onto microcapillaries of different materials under flow conditions using an enzyme assay.⁹² The adsorption of the enzyme onto the microcapillary surface was calculated based on the fractional enzyme loss. However, difficulties in resolving the resultant small changes in enzyme loss, especially at high initial enzyme concentrations, arose in this study. Determining the amount of adsorbed protein in a microchannel through a solution depletion method is therefore not a sensitive method of characterisation. Shortly after, Nicolau et al. modelled the adsorption of proteins onto microfluidic devices to estimate the thickness of the adsorbed protein layer at different protein concentrations, buffer ionic strengths and wall surface tensions.⁹⁵ Due to the complexities of the proteins, these studies were modelled based on a generic model protein derived from an average of 26 proteins in the online database.⁹⁷ Thus, these observations may not hold true for ‘non-generic’ proteins and surfaces with different characteristics. A laser induced fluorescence technique was used to probe protein adsorption onto microchannel surfaces by Lionello et al.^93,94 Immunoglobulin G was adsorbed onto a polymeric microchannel under both static and dynamic conditions. The group demonstrated adsorption kinetics coupled with diffusion phenomena in microchannels, and ways of loading the protein to obtain the best coverage for immunoassay purposes. Due to bulk solution depletion upon adsorption, multiple sample loads in ‘static’ conditions were found to be necessary to obtain an optimum surface coverage. Studies of fibrinogen adsorption on glass micro-capillaries have also been performed using an enzyme-linked immunoassay (ELISA) method, where the effects of protein adsorption in static versus dynamic adsorption conditions were investigated.⁵⁹ At any given time, the rate of adsorption in static conditions was always observed to be lower than the dynamic system due to bulk solution depletion. Adsorption kinetics of lysozyme on a hydrophobic cyclic olefin copolymer (COC) channel was recently studied by Alvarez, where the main focus was to demonstrate possibilities of real-time monitoring of protein adsorption using pulsed streaming potentials.⁸³ Although a non-labeling approach was used, a more complex design of the chip setup is required. Despite varying surface activities due to different experimental conditions from all these studies, it can be concluded that the nature of protein–surface interaction is not affected by miniaturisation; and that protein adsorbs rapidly onto the microchannel due to fast diffusion: the protein reaches the surface at a higher rate than its adsorption.

3.2 Surface modification in microchannels

The desire to control surface properties in many applications has led to intensive research on surface modification techniques. Surfaces with varying physical and chemical characteristics can subsequently be obtained, which can directly influence the biological performance at the interfaces.^98,99 In chip-CE for instance, higher protein separation efficiencies and migration time reproducibility were reported on coated-microchannels compared to non-coated microchannels due to the stable EOF and lower non-specific protein adsorption;¹⁰⁰ whereas higher peak capacities and separation resolution (narrow protein bands) were observed in chip-IEF.³⁰ Oriented antibodies on carefully controlled surfaces were also shown to improve the assay detection signal due to higher specific antigen capture efficiency.^101,102 The regulation of EOF and control of protein adsorption is therefore integral to enable rapid and efficient protein separation.^60,103,104

There are a number of surface modification strategies commonly used in biomedical and biotechnology applications, including wet chemical treatments such as self assembled monolayers, the Langmuir–Blodgett-Kuhn technique, polyelectrolyte multilayer deposition; and plasma modification.^105–109 In this review, we will primarily focus on surface modification techniques suitable for microchannels and specifically those used in the electrophoretic separation of biomolecules.

Surface modification of microchannels can be challenging, due to the range of materials utilised to create the devices. Common construction materials in microchip fabrication include glass, fused silica, quartz, silicon and polymeric materials such as poly(dimethylsiloxane) (PDMS), poly(carbonate), poly(methylmethacrylate) (PMMA), poly(ethylene terephthalate), poly(styrene), poly(propylene) and COC.^110–112 Therefore, a simple and versatile surface modification technique capable of coating a wide range of materials with excellent coating stability, desired control of EOF as well as biomolecules adsorption is of huge interest. This situation is further complicated by the requirement that coatings for electrophoresis and lab-on-chip applications are stable and retain their specific properties throughout their use.¹¹³ Surface modification in microchannels can generally be divided into two major coating categories, covalent or dynamic.^112–114

3.2.1 Dynamic coating. Dynamic coating methods are relatively simple, and require the addition of surface modifiers such as polymers or surfactants either to the running buffer (termed dynamic coating), or by rinsing the surface with modifier solutions prior to analyses (‘static’ physically adsorbed coating). These modifiers are physically adsorbed to the microchannel surfaces, usually by electrostatic interactions or H-bonding and van der Waals forces; and are most commonly used to alter EOF properties within microchannels.^{35,111,113–117}

Different types of polymeric coatings (neutral, anionic and cationic) can be physically adsorbed on microchannel walls. Neutral polymers such as polyethylene oxide (PEO), poly(saccharides), cellulose derivatives, poly(vinyl alcohol) (PVA), poly(acrylamide), poly(vinyl pyrrolidone) (PVP) have been widely used to suppress EOF and reduce the electrostatic interactions between analytes and microchannel walls.^113–118 Hydrophilic polymers such as PVA, poly(acrylamide), PEO, celluloses, and poly(saccharides) do not adsorb strongly on glass/fused silica surfaces, and the coatings can be easily removed.^116,117 The stabilities of physically adsorbed polymers are often reported to be pH dependent. For example, a physically adsorbed PEO coating has been reported to degrade at pH > 8, possibly due to competitive H-bonding among silanol group, hydroxyl and PEO.³² Suppression of EOF in PVA adsorbed surfaces was also reported to be not effective at pH > 4–8 due to its poor coating stability.¹¹⁹

Adsorption of charged polymers on surfaces changes the surface zeta potential and thus, the EOF. These polymers can therefore be used to control the magnitude and direction of the EOF. The most commonly used charged polymers are of cationic (positively charged) nature, enabling the separation of basic analytes and reversing the direction of EOF on hydroxyl-group rich bare glass or fused silica.¹¹⁶ Examples of cationic polymers include poly(brene), poly(arginine), poly(ethyleneimine) (PEI), poly(l-lysine) (PLL), and poly(allylamine hydrochloride) (PAH).^120–122 Anionic (negatively charged) polymers can be introduced on negatively charged surfaces via the assembly of polyelectrolyte multilayers, also known as layer-by-layer (LbL) assembly.^113,116,123LbL requires the deposition of alternately charged layers of polymer. This approach has been widely applied to coat glass, silica and oxidised PDMS microchannels since its introduction in 1991 by Decher et al.¹²⁴LbL of polyelectrolytes with an anionic layer as the final layer has been shown to reduce protein adsorption and provides a more stable EOF compared to bare glass or fused silica.¹¹¹ On uncoated surfaces, stable and reproducible EOF is difficult to achieve without excessive rinsing and etching pre-conditioning steps with strong NaOH and/or HCl solutions, which remove hydrocarbon contamination and regenerate surface OH groups.¹¹³ For example, polyelectrolyte multilayers with a final dextran sulfate layer has been used to separate proteins at (acidic) pH 2, which is not possible with uncoated glass (pI 2–3).¹²⁵

Other than the use of polymers as surface modifiers, the addition of small molecules such as surfactants, salts and various types of amine-containing molecules has also been investigated to reduce protein adsorption and changes the EOF behaviour of microchannels.^126–130Biopolymer coatings such as the ELISA blocking proteins, lysozyme, and fibrinogen have been utilised by several groups.^131–133 However, controlling the behaviour of these biopolymer coatings are always difficult, and specific and non-specific interactions between the coatings and the biomolecular species being separated may occur. This then leads to desorption, denaturation or exchange between the coating and the sample.

A wide range of surfactants are also used as surface modifiers within on-chip devices.^116,134,135 Anionic and cationic surfactants are often used to increase and decrease or even reverse the EOF properties within a channel. These surfactants however, tend to be strongly protein denaturing, and may not be suitable for analysis where maintaining the native conformation of proteins may be required.¹¹⁶ The use of zwitterionic surfactants such as 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS) has been shown to be able to neutralise the surface charge, therefore resulting in EOF suppression.¹³⁶ Despite the ability of nonionic surfactants to reduce EOF,^137,138 they have been generally shown to be much less effective in reducing the adsorption of proteins on surfaces compared to zwitterionic surfactants.¹¹⁵ Non-ionic surfactants generally have weaker interactions with proteins compared to ionic and zwitterionic surfactants.¹³⁹ The concentration of the surfactants used as surface modifiers must also be carefully manipulated to avoid the formation of micelles.

While physical adsorption can be readily applied to microfluidic and on chip devices, it has limitations. It poses environmental risks, due to the solvent chemical wastes.^74,111 Surfaces formed by physical adsorption are generally less stable and less reproducible, especially when they are reproduced on different types of substrates.^74,111 Different surface characteristics can also be generated from the same polymer due to different coating processes (polymer solutions, pretreatment solutions and coating protocols).^32,140Regeneration of the coatings is also often required with physical adsorption, as stabilities are often limited (usually after only a few EOF runs).¹³² This process is time consuming, and surface pretreatment steps usually require >30 min.³² The presence of surface additives in the running buffer (dynamic coating) or possible desorption of additives from the surfaces (‘static’ coating) may be detrimental towards applications requiring analysis by mass spectrometry and affinity CE due to possible ion suppression. Although being able to suppress EOF in many cases where separation resolution can be increased, these types of coatings do not effectively eliminate protein adsorption.^111,116

3.2.2 Covalent coating. Surface modification using covalent immobilisation is always preferred compared to physical or dynamic adsorption, due to the enhanced surface stability and the resulting higher resolution separations that can be achieved.¹¹⁴ Covalent immobilisation is often based on silane chemistries.¹⁴¹ The silane-coupling agent acts as an anchor between inorganic silanol containing substrates and the organic material coating. It consists of two steps: initial silane modification of the silanol surface groups (most commonly using bifunctional silanes), followed by incorporation of polymers or small molecules onto the silane surface. Silanisation processes can vary according to the different types of silane reagents, solvents, and surface pretreatment procedures, where the modified surfaces may vary accordingly.¹⁴² One of the main disadvantages of silanisation coating is the poor stability of the siloxane (Si–O–Si) bonds at high solution pH. Also, silanes are usually grafted on inorganic oxide containing surfaces such as glass and metal oxides; and it is often not applicable to polymeric materials.^98,111

Similar to dynamic coating, polymers such as poly(acrylamide), PVP, PVA, PEO, PEI, PLL, proteins, and other amine and acid containing species can be covalently bound to microchannel surfaces.^143–147 These can be coated using silane or non-silane based chemistry including radical polymerisation, thermal immobilisation and surface activation followed by solution based deposition.^{100,119,148–150} Covalently immobilised coatings have been reported to exhibit longer coating stability than physical adsorption. For example, covalent attachment of PEI on a PMMA surface exhibited a stable EOF of >50 runs;¹²² while >90 runs were observed for the corresponding PAH and PEI grafted glass surfaces (<60–90 runs for physically adsorbed polymers).³⁵

The covalent coating of polymers on surfaces requires reactive functional groups to be present at the surface to initiate immobilisation, and as such only certain materials can be used. The lack of a surface ionisable group in polymeric materials such as poly(ethylene), poly(propylene), PDMS, poly(carbonate) and poly(tetrafluoroethylene) (PTFE) requires surface pre-modification to introduce reactive functional groups.^151,152 This can be achieved using UV irradiation, laser activation, or plasma treatment.^{100,151,153,154} These treatments alone can be used to render hydrophobic surfaces hydrophilic, and can also result in an increase in EOF within channels formed in these polymeric materials due to the introduction of protonatable groups. For example, the EOF of plasma oxidised PDMS was reported to be 4 times higher than untreated PDMS.¹⁵⁵ An EOF increase of ca. 4% was observed on a PMMA laser treated surface.¹⁵⁴ It should, however, be noted that these surface pre-treatments are often dependent on the initial surface chemistry, subject to often rapid ageing effects including hydrophobic recovery and oxidation, and thus often show limited stability.^143,155 In short, many covalent coating methods for microfluidic devices require multi-step immobilisation procedures, which may be difficult to control, and can result in irreproducible coatings.¹¹⁷

3.2.3 Plasma-based coating

Commonly used solvent or liquid phase coatings may also cause problems, such as clogging or poor uniformities in the microchannels, so solventless deposition (plasma and non-plasma) has also been sought as an alternative method to coat microchannels.¹⁵⁶ Plasma surface modification can be divided into two classes, namely plasma treatment (chemically or physically altering the surface itself), and plasma polymerisation or deposition. Plasma treatments have been used to oxidise PDMS as well as other polymers, and enable the subsequent grafting of polymers or the bonding of devices together.^156,157 Plasma processing has also been shown to create multi-functional microfluidics with controllable surface wetting properties, allowing fluid manipulation, which is useful for directing protein and cell growth.¹⁵⁸ As detailed earlier, these treatments are generally effective for short periods of time.¹²⁶ Plasma polymerisation is a deposition rather than a treatment process that results in the formation of a polymeric thin film from the vapour phase under the influence of a plasma discharge.¹⁵⁹ Volatilised monomers are introduced into an evacuated reaction chamber (see Fig. 3) where, under the influence of an electric field, they undergo ionisation, generating electrons, ions, free radicals, photons, and molecules both in ground and excited states. Excitation of the monomer results in reactive species on the surface, which act as reactive sites for the covalent attachment of other species.^160,161 It can therefore be viewed as a kind of graft polymerisation, although it does not alter the bulk properties of the substrate by partial penetration of the substrate, as is the case in graft polymerisation obtained by conventional means.¹⁵⁹ Plasma processing is a complex process with repetitive fragmentation, rearrangement, crosslinking, ionisation, and polymerisation of the monomers occurring throughout the cycle. The resulting polymer thin film is deposited on the surfaces of any material exposed to the plasma glow. Plasma polymers have been utilised in biosensors,¹⁶² surface micropatterning,¹⁶³ blood contacting devices,¹⁶⁴cell culture surfaces,¹⁶⁵ biomolecule immobilisation,¹⁶⁶ and polymer grafting.¹⁶⁰

Fig. 3 Schematic example of a radio frequency (rf) plasma polymer reactor [Reproduced from supplementary material in ref. 35].

The polymers formed by plasma polymerisation tend to be highly branched and cross-linked with random structures incorporating various structural fragments of the monomer.^159,161 However, by controlling the plasma deposition parameters, it is increasingly possible to produce more traditional polymer structures, chemistries and behaviours.^167–169 Plasma polymers have a number of advantages over other surface modification techniques: good adhesion and ability to coat virtually any dry substrate; the coatings do not affect the bulk properties of the substrate; a solvent-free and one-step coating process; ultra-thin films (1–10 nm) with controllable thickness; a large number of different monomers have been shown to form films.^159,160,170

In 2003, the Karube group investigated the first use of a plasma polymerisation to coat glass open-microchannels for IEF-based protein separation.^171,172Hexamethyldisiloxane (HMDSO), glycidol, and acetonitrile were used as the monomers to reduce EOF. It was reported that the plasma polymerised acetonitrile coating was easily removed due to weak surface attachment; and a minimum thickness of 50 nm had to be obtained for HMDSO and glycidol coatings to remain stable at high applied voltages. Barbier et al. introduced plasma polymerised acrylic acid on PDMS microchannels and these exhibited stability over several days.¹⁷³ Coatings produced on PDMS have been widely known to be susceptible to degradation due to fast hydrophobic recovery.¹⁷⁴ General surface modification strategies and their recent development for PDMS microfluidic chips have been reviewed by several groups.^175,176 It was suggested that a combination of both wet chemical treatments and solventless techniques provided the greatest promise in terms of surface stability. Incorporating additives such as carboxylic acid and PEG-based polymers to the PDMS pre-polymer prior to curing has also been shown to improve surface wettability and provide better control of EOF as well as non-specific protein adsorption.^177,178 A stable and non-fouling coating was then demonstrated by Salim et al. in glass microchannels using a tetraglyme plasma polymer.¹⁷⁹ This work has recently been adapted to produce coatings on the more challenging PDMS substrates that are stable and resist protein adsorption for up to 100 days [A. G. Pereira-Medranoet al. Rapid fabrication of Glass/PDMS hybrid μIMER for augmented high throughput membrane proteomics, submitted to Lab Chip 2010].

These early studies suggest plasma polymerisation as a versatile method to coat microchannel surfaces. To date, wet chemistries are still the preferred choice to surface modify microchannels, as solutions can be directly flowed through the ‘closed’ microfluidic chip system. Coating of the inner microchannel surfaces using plasma can be challenging; and an ‘open-system’ is usually devised, where the plasma-coated channel has to be subsequently bonded to another cover to form a seal. Consequently, miniaturised plasma sources/microplasma is being increasingly investigated to coat inner surface of microchannels and small tubings.^180–182 Although several groups have shown the successful use of microplasma to produce thin film coatings to control EOF and biomolecule adsorption,^183,184 this technology is still in its early development phase. Design improvements and understanding of the microplasma properties are essential to obtain controlled, uniform film properties.

4. Understanding coatings that resist and bind biomolecules

4.1 Surfaces that resist bioadhesion

Various coatings such as PEO, phospholipids, poly(saccharides), poly(acrylamide), and proteins have been presented for their ability to reduce bioadhesion in a range of biomedical and biotechnology applications.^185–192 Of these, PEO was shown to offer great promise, and has been widely investigated. The structure of PEO is HO–(CH₂–O–CH₂)_n–OH,¹⁹³ where n may range from 0–100 000.¹⁹⁴PEO-based surfaces can be used to create a non-fouling platform suitable in applications where elimination of protein adsorption is essential. By using a mixture of reactive and non-reactive grafted PEG chains, it is also possible to immobilise specific proteins and other biomolecules to a non-fouling PEG surface. PEO surfaces can be produced by several approaches, including physical adsorption,^32,195 SAMs,^196,197 covalent immobilisation,^198,199 vapour deposition techniques,¹⁵⁶ and plasma polymerisation of ethylene oxide monomers.^{163,188,200–205}

In aqueous solutions, the PEO molecule is hydrophilic and highly mobile with a large exclusion volume.¹⁹³ It has a strong tendency to form hydrogen bonds with water, and this extensive hydration of PEO molecule causes the van der Waals attraction to be small and no electrostatic attraction occurs.²⁰⁶ The highly mobile or rapid motion of the PEO molecule is also reported to give the approaching protein little time to form an interaction.²⁰⁶ The mechanisms for the antifouling properties of PEO are not yet fully understood, and are often related to the chain length and surface density of PEO, which gives rise to steric repulsion effects.^207–210 Steric repulsion effects (osmotic and excluded volume contributions) arise from the compression of the PEO chains, where there is a loss in entropy as a protein approaches the surfaces. Adsorption of a protein onto the PEO surface is therefore thermodynamically unfavourable.^74,206

Jeon et al. reported a balance between steric repulsion, van der Waals and hydrophobic interactions between protein molecules and PEO surfaces, where the net force which determines the adsorption of protein onto the PEO surface depends on the thickness of the PEO layers as well as the surface coverage.²⁰⁷ Sufficient layer thickness and polymer chain density can screen the protein-substrate interaction, and prevent diffusion of the protein through the steric layer to the underlying substrate.^209,210 Szleifer et al. reported that surface density has a greater effect in resisting protein adsorption than chain length.²¹¹ Overall, PEO with molecular weights <10 000 Da, and high chain density are generally reported to be effective in resisting protein adsorption.^151,210 High density polymers are, however, difficult to obtain with covalent attachment due to the steric exclusion volume of the attached polymer chain, which then hinders the attachment of another polymer chain.

In the radio frequency glow discharge (rfgd) plasma deposition process however, short EO containing molecules (tetraglyme, triglyme) can also produce surfaces with effective non-fouling properties. This may consist of a carpet of short length EO units –(CH₂–CH₂–O)_n– packed in a high density manner with a reduced steric exclusion volume.^200,202,204 Monomers of up to two EO units (diglyme) have also been shown to effectively resist protein adsorption.²⁰⁵ Long PEO chain units are therefore not necessary in rendering a surface non-fouling. Plasma polymerised surfaces of EO units are therefore postulated to be similar to the surfaces formed by the SAMs of EO terminated alkanethiols on gold.^203,204 It was first demonstrated in 1991 by Prime and Whitesides, that SAMs of short chain PEO can exhibit low bioadhesion.²¹² The surfaces formed by SAMs, however, are dependent on the substrate materials, with noble metal surfaces (Au, Ag) required to form the alkanethiol bond.^213,214

Although still under widespread investigation, the non-fouling properties of EO-based plasma polymerised surfaces are generally being correlated with the high retention of the C–O units within the plasma polymer structure. High retention of the C–O content can be achieved by lowering the plasma deposition power. By adjusting the plasma deposition power, different types of surface chemistries (i.e. extents of C–O incorporation) can be obtained from a single monomer system, producing films with varying protein adhesion properties.^204,215 Based on the data from the SAM literature, it is thought that the strong tendencies for the EO units to form hydrogen bonds with water may effectively screen or form a physical barrier to the surfaces from biomolecules.^205,216 Studies conducted by Shen et al. using ESCA and ToF-SIMS showed the presence of methyl-terminated short EO chain ends in the plasma deposited tetraglyme surface.²⁰³ The plasma polymerised surface formed however, may be more complex and may have disordered molecular surface structures as opposed to ordered SAMs.²⁰³ The EO units formed in the plasma polymer films probably do not have high mobilities or the conformational freedom as normally associated with grafted polymers as densely packed, short chain films are formed.²⁰² This densely packed complete surface coverage is thought to be a factor in the ability of the coatings to resist protein adsorption.²¹⁷ Therefore, conformational flexibility may not contribute to the protein resistivity in this case.²¹⁸

Plasma polymerised tetraglyme surfaces have been shown to able to resist fibrinogen adsorption up to a level of lower than 10 ng/cm², and thus resist cell adhesion.^179,203 Although plasma polymerisation is capable of producing ultrathin films (1–10 nm), there also appears to be a threshold of film thickness in the plasma polymerisation process whereby an effective non-fouling surface can be obtained. Chu et al. found a minimum film thickness of about 3 nm and 6 nm for resisting BSA and fibrinogen adsorption respectively on a plasma polymerised diglyme surface.²⁰¹ However, the exact surface chemistries for the different film thicknesses formed remained unclear, and may be due to a number of factors including the coverage of the plasma film and specific characteristics of plasma deposition system used.²⁰¹

4.2 Surfaces for functional immobilisation of biomolecules

Proteins contain functional groups such as –COOH, –NH₂, –OH and –SH, which can be physically (random) or covalently (oriented) immobilised to microchannel surfaces (Fig. 4).²¹⁹ In some lab-on-chip applications such as affinity separation^220,221 and enzymatic reactors,^222,223 biomolecules are required to be surface-immobilised in specific orientations such that their biological activities are retained. Reactive surface chemistries commonly used for biomolecule immobilisation are amine, carboxyl, aldehyde, epoxy, isocyanate, hydroxyl, maleimide, and NHS-activated surfaces, which can be prepared by different strategies: generally wet chemical treatments.⁷⁴ However, hydroxyl (alcohol monomers), amine (alkylamine monomers), carboxyl (carboxylic acid, propanoic acid monomer) and aldehyde (aldehye monomers) functionalised plasma polymers have also been shown to be successful for covalent immobilisation of biologically active molecules.²²⁴

Fig. 4 Schematic diagram of random, uncontrolled (A) and oriented (B) protein adsorption to surfaces.

5. Closing remarks

The central concept of the proteomics-on-chip is to integrate different types of separation and preparation tools. The aim of this is to enable fast analyses of a real-time complex sample containing up to thousands of proteins and many hundreds of thousands of peptides, present with possibility >10 orders (or even more) of magnitude in concentration range.²²⁵ Design of a universal microfluidic platform for proteomics is therefore difficult to predict due to the inherent complexities in each of the relevant unit operations, and many possible ways of integration. Most of the microfluidic chip systems built is therefore focused on specific applications such as on-line sample pre-treatment, protein digestion, protein/peptide separation, and its MS coupling. Regardless of the design, biomolecules that are present in most of the unit operations need to be effectively controlled by surface modification to obtain a globally reliable, robust, and reproducible microfluidic chip platform for complex proteome analyses. Although significant contributions have been made to control bio-interfacial interactions as already described in this article, further work is needed to effectively coat the inner surface of microchannels in a manner independent of the substrate materials used, and to develop characterisation tools for inner surface coatings and protein coverage. Integration between unit operations is also a major drawback that needs to be assessed to ensure down-stream solvent compatibility with MS instrumentation and minimal sample loss (so that low abundance proteins are still possible to measure). Addressing these challenges, together with the advancement in proteomics, microfluidics, and surface technologies, can provide the key to future growth in biotechnology, pharmaceutical, and biomedical research.

Acknowledgements

We acknowledge the EPSRC for funding (grants GR/S84347/01 and EP/E036252/1).

References

1. W. C. Andrea, Lab-on-a-chip: Opportunities for chemical engineering, AIChE J., 2002, 48(8), 1590–1595.
2. A. Manz, N. Graber and H. M. Widmer, Miniaturized total chemical analysis systems: A novel concept for chemical sensing, Sens. Actuators, B, 1990, 1(1–6), 244–248.
3. D. Janasek, J. Franzke and A. Manz, Scaling and the design of miniaturized chemical-analysis systems, Nature, 2006, 442(7101), 374–380.
4. N. Lion, F. Reymond, H. H. Girault and J. S. Rossier, Why the move to microfluidics for protein analysis?, Curr. Opin. Biotechnol., 2004, 15(1), 31–37.
5. U. Bilitewski, M. Genrich, S. Kadow and G. Mersal, Biochemical analysis with microfluidic systems, Anal. Bioanal. Chem., 2003, 377(3), 556–569.
6. M. J. Madou and R. Cubicciotti, Scaling issues in chemical and biological sensors, Proc. IEEE, 2003, 91(6), 830–838.
7. J. S. Rossier and H. H. Girault, Enzyme linked immunosorbent assay on a microchip with electrochemical detection, Lab Chip, 2001, 1(2), 153–157.
8. J. S. Rossier, G. Gokulrangan, H. H. Girault, S. Svojanovsky and G. S. Wilson, Characterization of Protein Adsorption and Immunosorption Kinetics in Photoablated Polymer Microchannels, Langmuir, 2000, 16(22), 8489–8494.
9. P. S. Dittrich and A. Manz, Lab-on-a-chip: microfluidics in drug discovery, 2006, 5(3), 210–218.
10. Garcia, D. Carlos, L. I. U. Yan, Henry and S. Charles, Recent progress in micro-TAS for clinical applications, Analyst, 2003, 128, 1002–1008.
11. C. H. Arnaud, Mimicking Biological Systems, Chemical and Engineering News, 2007, 14–19.
12. K.-J. Jang and K.-Y. Suh, A multi-layer microfluidic device for efficient culture and analysis of renal tubular cells, Lab Chip, 2010, 10, 36,  10.1039/b907515a.
13. D. Huh, A. Mammoto, M. Montoya-Zavala and D. E. Ingber, A human breathing lung-on-a-chip, FASEB J., 2010, 24 , (1_MeetingAbstracts), 1065.22-.
14. H. Yu, I. Meyvantsson, I. A. Shkel and D. J. Beebe, Diffusion dependent cell behavior in microenvironments, Lab Chip, 2005, 5(10), 1089–1095.
15. J. Roboz, Strategies, Techniques, and Applications in Cancer Biochemistry and Biology, CRC Press, 2002, p. 361.
16. R. E. Banks, M. J. Dunn, D. F. Hochstrasser, J.-C. Sanchez, W. Blackstock, D. J. Pappin and P. J. Selby, Proteomics: new perspectives, new biomedical opportunities, Lancet, 2000, 356(9243), 1749–1756.
17. P. Jagroop, P. L. S. Ambrosius, C. W. Phillip and A. B. Catherine, A cross-species quantitative proteomic study of salt adaptation in a halotolerant environmental isolate using 15N metabolic labelling, Proteomics, 2008, 8(11), 2266–2284.
18. J. Khandurina and A. Guttman, Bioanalysis in microfluidic devices, J. Chromatogr., A, 2002, 943(2), 159–83.
19. R. B. M. Schasfoort, Proteomics-on-a-chip: the challenge to couple lab-on-a-chip unit operations, Expert Rev. Proteomics, 2004, 1(1), 123–132.
20. D. Figeys and D. Pinto, Proteomics on a chip: promising developments, Electrophoresis, 2001, 22(2), 208–216.
21. S. Mouradian, Lab-on-a-chip: applications in proteomics, Curr. Opin. Chem. Biol., 2002, 6(1), 51–56.
22. J. Han and A. K. Singh, Rapid protein separations in ultra-short microchannels: microchip sodium dodecyl sulfate-polyacrylamide gel electrophoresis and isoelectric focusing, Journal of chromatography. A, 2004, 1049(1–2), 205–209.
23. Y. Li, J. S. Buch, F. Rosenberger, D. L. DeVoe and C. S. Lee, Integration of isoelectric focusing with parallel sodium dodecyl sulfate gel electrophoresis for multidimensional protein separations in a plastic microfluidic [correction of microfludic] network, Anal. Chem., 2004, 76(3), 742–748.
24. J. Kim, M. Johnson, P. Hill and B. K. Gale, Microfluidic sample preparation: cell lysis and nucleic acid purification, Integr. Biol., 2009, 1(10), 574–586.
25. Y. Hiroshi, M. Masaya, H. Takeshi, B.-N. Maria Portia, A. Kazunari and M. Hideaki, Rapid and efficient proteolysis for proteomic analysis by protease-immobilized microreactor, Electrophoresis, 2009, 30(18), 3257–3264.
26. J. Carlier, K. Chuda, S. Arscott, V. Thomy, B. Verbeke, X. Coqueret, C. Jean Christophe, C. Druon and P. Tabourier, High pressure-resistant SU-8 microchannels for monolithic porous structure integration, J. Micromech. Microeng., 2006, 16, 2211–2219.
27. L. Jeonghoon, A. S. Steven and K. M. Kermit, Microfluidic chips for mass spectrometry-based proteomics, J. Mass Spectrom., 2009, 44(5), 579–593.
28. J. Lee, S. A. Soper and K. K. Murray, Microfluidics with MALDI analysis for proteomics—A review, Anal. Chim. Acta, 2009, 649(2), 180–190.
29. R. T. Kelly, K. Tang, D. Irimia, M. Toner and R. D. Smith, Elastomeric Microchip Electrospray Emitter for Stable Cone-Jet Mode Operation in the Nanoflow Regime, Anal. Chem., 2008, 80(10), 3824–3831.
30. C. Li, Y. Yang, H. G. Craighead and K. H. Lee, Isoelectric focusing in cyclic olefin copolymer microfluidic channels coated by polyacrylamide using a UV photografting method, Electrophoresis, 2005, 26(9), 1800–1806.
31. H.-B. Javier, N. Christian, C. Alejandro and P. Matthias, On-line capillary electrophoresis-mass spectrometry for the analysis of biomolecules, Electrophoresis, 2004, 25(14), 2257–2281.
32. J. Preisler and E. S. Yeung, Characterization of Nonbonded Poly(ethylene oxide) Coating for Capillary Electrophoresis via Continuous Monitoring of Electroosmotic Flow, Anal. Chem., 1996, 68(17), 2885–2889.
33. G. Y. Tang, C. Yang, J. C. Chai and H. Q. Gong, Joule heating effect on electroosmotic flow and mass species transport in a microcapillary, Int. J. Heat Mass Transfer, 2004, 47(2), 215–227.
34. S. Ghosal, Effect of Analyte Adsorption on the Electroosmotic Flow in Microfluidic Channels, Anal. Chem., 2002, 74(4), 771–775.
35. M. Salim, P. C. Wright and S. L. McArthur, Studies of electroosmotic flow and the effects of protein adsorption in plasma-polymerized microchannel surfaces, Electrophoresis, 2009, 30(11), 1877–1887.
36. W. Hellmich, J. Regtmeier, T. T. Duong, R. Ros, D. Anselmetti and A. Ros, Poly(oxyethylene) Based Surface Coatings for Poly(dimethylsiloxane) Microchannels, Langmuir, 2005, 21(16), 7551–7557.
37. R. E. Oosterbroek and A. v. d. Berg, Lab-on-a-chip: miniaturized systems for (bio) chemical analysis and synthesis, Elsevier B.V, 2003, p. 450.
38. A. J. de Mello and N. Beard, Dealing with real samples: sample pre-treatment in microfluidic systems, Lab Chip, 2003, 3(1), 11N–19.
39. Y.-C. Tyan, J.-D. Liao, S.-B. Jong, P.-C. Liao, M.-H. Yang, Y.-W. Chang, R. Klauser, M. Himmelhaus and M. Grunze, Characterization of trypsin immobilized on the functionable alkylthiolate self-assembled monolayers: a preliminary application for trypsin digestion chip on protein identification using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry, J. Mater. Sci.: Mater. Med., 2005, 16(2), 135–142.
40. A. Ríos, A. Escarpa, M. C. González and A. G. Crevillén, Challenges of analytical microsystems, TrAC, Trends Anal. Chem., 2006, 25(5), 467–479.
41. D. Mark, S. Haeberle, G. Roth, F. v. Stetten and R. Zengerle, Microfluidic lab-on-a-chip platforms: requirements, characteristics and applications, Chem. Soc. Rev., 2010, 39(3), 1153–1182.
42. T. Nguyet Thuy, A. Ichraf, P. Antoine and T. Myriam, Recent innovations in protein separation on microchips by electrophoretic methods: An update, Electrophoresis, 2010, 31(1), 147–173.
43. W. Norde, Adsorption of proteins from solution at the solid–liquid interface, Adv. Colloid Interface Sci., 1986, 25, 267–340.
44. J. D. Andrade, Surface and Interfacial Aspects of Bio-medical Polymers, in Principles of protein adsorption, Plenum Press, New York, 1985, vol. 2, pp. 4–6.
45. C. A. Haynes and W. Norde, Globular proteins at solid/liquid interfaces, Colloids Surf., B, 1994, 2(6), 517–566.
46. J. D. Andrade, V. Hlady and A. P. Wei, Adsorption of complex proteins at interfaces, Pure Appl. Chem., 1992, 64(11), 1777–1781.
47. K. Al-Malah, in Adsorption: Theory, Modeling, and Analysis, ed. J. Tóth, Marcel Dekker, New York, 2002, vol. 107, p. 803.
48. P. M. Claesson, E. Blomberg, J. C. Froberg, T. Nylander and T. Arnebrant, Protein interactions at solid surfaces, Adv. Colloid Interface Sci., 1995, 57, 161–227.
49. K. C. Dee, R. Bizios and D. A. Puleo, Protein-surface Interactions, in Introduction to Tissue-Biomaterial Interactions, John Wiley and Sons Ltd., 1st edn, 2002, pp. 37–40.
50. C. F. Wertz and M. M. Santore, Effect of Surface Hydrophobicity on Adsorption and Relaxation Kinetics of Albumin and Fibrinogen: Single-Species and Competitive Behavior, Langmuir, 2001, 17(10), 3006–3016.
51. M. van der Veen, M. C. Stuart and W. Norde, Spreading of proteins and its effect on adsorption and desorption kinetics, Colloids Surf., B, 2007, 54(2), 136–142.
52. J. Buijs and V. Hlady, Adsorption Kinetics, Conformation, and Mobility of the Growth Hormone and Lysozyme on Solid Surfaces, Studied with TIRF, J. Colloid Interface Sci., 1997, 190(1), 171–181.
53. M. Malmsten, Ellipsometry studies of the effects of surface hydrophobicity on protein adsorption, Colloids Surf., B, 1995, 3(5), 297–308.
54. G. B. Sigal, M. Mrksich and G. M. Whitesides, Effect of Surface Wettability on the Adsorption of Proteins and Detergents, J. Am. Chem. Soc., 1998, 120(14), 3464–3473.
55. T. Zoungrana, G. H. Findenegg and W. Norde, Structure, Stability, and Activity of Adsorbed Enzymes, J. Colloid Interface Sci., 1997, 190(2), 437–448.
56. G. Raffaini and F. Ganazzoli, Protein Adsorption on a Hydrophobic Surface: A Molecular Dynamics Study of Lysozyme on Graphite, Langmuir, 2010, 26(8), 5679–5689.
57. Z. Adamczyk, Irreversible adsorption of particles, in Adsorption: Theory, Modeling, and Analysis, ed. J. Tóth, Marcel Dekker, 2002, vol. 107, p. 251.
58. S. Kumar and R. Nussinov, Close-Range Electrostatic Interactions in Proteins, ChemBioChem, 2002, 3(7), 604–617.
59. M. Salim, B. O'Sullivan, S. L. McArthur and P. C. Wright, Characterization of fibrinogen adsorption onto glass microcapillary surfaces by ELISA, Lab Chip, 2007, 7(1), 64–70.
60. R. A. Latour, Biomaterials: Protein–Surface Interactions, in Encyclopedia of Biomaterials and Biomedical Engineering, ed. Gary L. Bowlin and G. W., Marcel Dekker, Inc., New York, 2005, vol. 2, p. 1.
61. L. Hagel, Separation on the basis of chemistry, in Protein Purification Techniques: A Practical Approach, ed. S. Roe, Oxford University Press, 2nd edn, 2001, p. 155.
62. K. L. Hanson, L. Filipponi and D. V. Nicolau, Biomolecules and cells on surfaces-fundamental concepts, in Microarray Technology and its Application, ed. U. R. Müller and D. V. Nicolau, Springer, New York, 2005, pp. 23–44.
63. A. T. Hubbard, Encyclopedia of Surface and Colloid Science, Marcel Dekker, New York, 2002, vol. 4, p. 4383.
64. T. Arakawa and J. S. Philo, Biophysical and biochemical analysis of recombinant proteins structure and analysis of proteins, in Pharmaceutical Biotechnology, D. J. A. Crommelin and R. D. Sindelar, Gutenberg Press, 2nd edn, 2002, p. 25.
65. J. L. Brash and T. A. Horbett, Proteins at Interfaces: Physicochemical and Biochemical Studies, Washington DC, 1987, ACS Symposium Series, vol. 343, pp. 1–33.
66. J. L. Brash and T. A. Horbett, Proteins at interfaces II: fundamentals and applications, Washington DC, 1995, ACS Symposium Series, vol. 602, pp. 1–23.
67. J. D. Andrade, V. Hlady, A.-P. Wei, C.-H. Ho, A. S. Lea, S. I. Jeon, Y. S. Lin and E. Stroup, Proteins at interfaces: Principles, multivariate aspects, protein resistant surfaces, and direct imaging and…, Clin. Mater., 1992, 11(1–4), 67–84.
68. M. Malmsten, Protein Adsorption at the Solid–Liquid Interface, in Protein Architecture, ed. Y. Lvov and H. Hohwald, Marcel Dekker, 1999, p. 6.
69. Y. Ling, Z. Lu, Y. Gan, Y. Liu and C. M. Li, AFM study of adsorption of protein A on a poly(dimethylsiloxane) surface, Nanotechnology, 2009, 20(28), 285101.
70. E. Ouellet, C. Lausted, T. Lin, C. W. T. Yang, L. Hood and E. T. Lagally, Parallel microfluidic surface plasmon resonance imaging arrays, Lab Chip, 2010, 10(5), 581–588.
71. G. Krishnamoorthy, E. T. Carlen, H. L. deBoer, A. van den Berg and R. B. M. Schasfoort, Electrokinetic Lab-on-a-BioChip for Multi-ligand/Multi-analyte Biosensing, Anal. Chem., 2010, 82(10), 4145–4150.
72. J. S. M. Hans Griesser, S. Matthew Wagner, G. David Castner, Peter Kingshott and M. Keith Mclean, XPS, ToF-SIMS, and MALDI-MS for Characterizing Adsorbed Protein Films, in Biopolymers at Interfaces, ed. M. Malmsten, CRC Press, 2003, vol. 110.
73. H. H. Peter Kingshott, Methods of assessing protein adsorption, in Encyclopedia of Surface and Colloid Science, ed. T. A. Hubbard, CRC Press, 2002, vol. 3, p. 3343.
74. J. M. Goddard and J. H. Hotchkiss, Polymer surface modification for the attachment of bioactive compounds, Prog. Polym. Sci., 2007, 32(7), 698–725.
75. S. L. McGurk, R. J. Green, G. H. W. Sanders, M. C. Davies, C. J. Roberts, S. J. B. Tendler and P. M. Williams, Molecular interactions of biomolecules with surface-engineered interfaces using atomic force microscopy and surface plasmon resonance, Langmuir, 1999, 15(15), 5136–5140.
76. Y. F. Dufrene, T. G. Marchal and P. G. Rouxhet, Probing the organization of adsorbed protein layers: Complementarity of atomic force microscopy, X-ray photoelectron spectroscopy and radiolabeling, Appl. Surf. Sci., 1999, 144–145, 638–643.
77. M. Bergkvist, Adsorption of Proteins: Methods of Orientation, in Encyclopedia of Surface and Colloid Science, ed. P. Somasundaran, CRC Press, 2006, vol. 1.
78. D. S. Mantus, B. D. Ratner, B. A. Carlson and J. F. Moulder, Static secondary ion mass spectrometry of adsorbed proteins, Anal. Chem., 1993, 65(10), 1431–1438.
79. J. Marchand-Brynaert, Polymer Membranes, in Encyclopedia of Surface and Colloid Science, ed. P. Somasundaran, CRC Press, 2006, vol. 6, p. 4861.
80. H. Matsumura, K. Kawasaki, N. Okumura, M. Kambara and W. Norde, Characterization of the surface of protein-adsorbed dental materials by wetting and streaming potential measurements, Colloids Surf., B, 2003, 32(2), 97–103.
81. J. W. Pollard, Two-Dimensional Polyacrylamide Gel Electrophoresis of Proteins, in Methods in Molecular Biology, ed. J. M. Walker, Humana Press Inc., New Jersey, 1984, vol. 1, p. 81.
82. W. Possart, Surface Characterization of Advanced Polymers, ed. Luigia Sabbatini and Pier Giorgio, Zambonin VCH, Weinheim, New York, Basel, Cambridge, Tokyo, 1993. Acta Polymerica, 1994, 45, (2), 131.
83. F. Luna-Vera and J. C. Alvarez, Adsorption kinetics of proteins in plastic microfluidic channels: Real-time monitoring of lysozyme adsorption by pulsed streaming potentials, Biosens. Bioelectron., 2010, 25(6), 1539–1543.
84. F. Boumaza, P. Dejardin, F. Yan, F. Bauduin and Y. Holl, Fibrinogen adsorption on Pyrex glass tubes: A continuous kinetic study, Biophys. Chem., 1992, 42(1), 87–92.
85. J. N. B. D. Lin, A. S. Lea, P. K. Hansma and J. D. Andrade, Direct observation of Immunoglobulin adsorption dynamics using the atomic force microscope, Langmuir, 1990, 6, 509–511.
86. Y.-S. Lin and V. Hlady, Human serum albumin adsorption onto octadecyldimethylsilyl–silica gradient surface, Colloids Surf., B, 1994, 2(5), 481–491.
87. J. L. Ortega-Vinuesa, P. Tengvall and I. Lundstrom, Molecular packing of HSA, IgG, and fibrinogen adsorbed on silicon by AFM imaging, Thin Solid Films, 1998, 324(1–2), 257–273.
88. W. J. J. Dillman and I. F. Miller, On the adsorption of serum proteins on polymer membrane surfaces, J. Colloid Interface Sci., 1973, 44(2), 221–241.
89. L. B. Thompson, N. H. Mack and R. G. Nuzzo, Bifunctional polyacrylamide based polymers for the specific binding of hexahistidine tagged proteins on gold surfaces, Phys. Chem. Chem. Phys., 2010, 12(17), 4301–4308.
90. N. Bonnet, D. O'Hagan and G. Hahner, Protein adsorption onto CF3-terminated oligo(ethylene glycol) containing self-assembled monolayers (SAMs): the influence of ionic strength and electrostatic forces, Phys. Chem. Chem. Phys., 2010, 12(17), 4367–4374.
91. J. Jenkins, B. Prabhakarpandian, K. Lenghaus, J. Hickman and S. Sundaram, Fluidics-resolved estimation of protein adsorption kinetics in a biomicrofluidic system, Anal. Biochem., 2004, 331(2), 207–215.
92. K. Lenghaus, J. W. Dale, J. C. Henderson, D. C. Henry, E. R. Loghin and J. J. Hickman, Enzymes as Ultrasensitive Probes for Protein Adsorption in Flow Systems, Langmuir, 2003, 19(15), 5971–5974.
93. A. Lionello, J. Josserand, H. Jensen and H. H. Girault, Dynamic protein adsorption in microchannels by “stop-flow” and continuous flow, Lab Chip, 2005, 5(10), 1096–103.
94. A. Lionello, J. Josserand, H. Jensen and H. H. Girault, Protein adsorption in static microsystems: effect of the surface to volume ratio, Lab Chip, 2005, 5(3), 254–60.
95. D. V. Nicolau, Jr., F. Fulga and D. V. Nicolau, Impact of protein adsorption on the geometry of microfluidic devices, Biomed. Microdevices, 2003, 5(3), 227–233.
96. W. F. Patton, Detection technologies in proteome analysis, J. Chromatogr., B: Anal. Technol. Biomed. Life Sci., 2002, 771(1–2), 3–31.
97. E. N. Vasina, E. Paszek, D. V. Nicolau, Jr. and D. V. Nicolau, The BAD project: data mining, database and prediction of protein adsorption on surfaces, Lab Chip, 2009, 9(7), 891–900.
98. N. Nath, J. Hyun, H. Ma and A. Chilkoti, Surface engineering strategies for control of protein and cell interactions, Surf. Sci., 2004, 570(1–2), 98–110.
99. M. Mrksich, What can surface chemistry do for cell biology?, Curr. Opin. Chem. Biol., 2002, 6(6), 794–797.
100. J. Liu, T. Pan, A. T. Woolley and M. L. Lee, Surface-Modified Poly(methyl methacrylate) Capillary Electrophoresis Microchips for Protein and Peptide Analysis, Anal. Chem., 2004, 76(23), 6948–6955.
101. Y. Yuan, H. Hongyan and L. J. Lee, Protein A-based antibody immobilization onto polymeric microdevices for enhanced sensitivity of enzyme-linked immunosorbent assay, Biotechnol. Bioeng., 2009, 102(3), 891–901.
102. W.-C. Sung, H.-H. Chen, H. Makamba and S.-H. Chen, Functionalized 3D-Hydrogel Plugs Covalently Patterned Inside Hydrophilic Poly(dimethylsiloxane) Microchannels for Flow-Through Immunoassays, Anal. Chem., 2009, 81(19), 7967–7973.
103. D. G. Castner and B. D. Ratner, Biomedical surface science: Foundations to frontiers, Surf. Sci., 2002, 500(1–3), 28–60.
104. W. L. W. Hau, D. W. Trau, N. J. Sucher, M. Wong and Y. Zohar, Surface-chemistry technology for microfluidics, J. Micromech. Microeng., 2003, 13(2), 272.
105. M. Tirrell, Chemical engineering of polymers: production of flexible, functional materials, Chem. Eng. Sci., 1995, 50(24), 4123–4141.
106. M. Advincula, X. Fan, J. Lemons and R. Advincula, Surface modification of surface sol–gel derived titanium oxide films by self-assembled monolayers (SAMs) and non-specific protein adsorption studies, Colloids Surf., B, 2005, 42(1), 29–43.
107. M. Herrero, R. Navarro, Y. Grohens, H. Reinecke and C. Mijangos, Controlled wet-chemical modification and bacterial adhesion on PVC-surfaces, Polym. Degrad. Stab., 2006, 91(9), 1915–1918.
108. C. Oehr, Plasma surface modification of polymers for biomedical use, Nucl. Instrum. Methods Phys. Res., Sect. B, 2003, 208, 40–47.
109. Y. L. Lee and C. Y. Chen, Surface wettability and platelet adhesion studies on Langmuir–Blodgett films, Appl. Surf. Sci., 2003, 207(1–4), 51–62.
110. K. Huikko, R. Kostiainen and T. Kotiaho, Introduction to micro-analytical systems: bioanalytical and pharmaceutical applications, Eur. J. Pharm. Sci., 2003, 20(2), 149–71.
111. A. Pallandre, B. de Lambert, R. Attia, A. M. Jonas and J.-L. Viovy, Surface treatment and characterization: perspectives to electrophoresis and lab-on-chips, Electrophoresis, 2006, 27(3), 584–610.
112. J. Liu and M. L. Lee, Permanent surface modification of polymeric capillary electrophoresis microchips for protein and peptide analysis, Electrophoresis, 2006, 27(18), 3533–3546.
113. D. Belder and M. Ludwig, Surface modification in microchip electrophoresis, Electrophoresis, 2003, 24(21), 3595–606.
114. V. Dolník, Wall coating for capillary electrophoresis on microchips, Electrophoresis, 2004, 25(21–22), 3589–3601.
115. P. G. Righetti, C. Gelfi, B. Verzola and L. Castelletti, The state of the art of dynamic coatings, Electrophoresis, 2001, 22(4), 603–611.
116. E. A. S. Doherty, R. J. Meagher, M. N. Albarghouthi and A. E. Barron, Microchannel wall coatings for protein separations by capillary and chip electrophoresis, Electrophoresis, 2003, 24(12), 34–54.
117. J. Horvath and V. Dolník, Polymer wall coatings for capillary electrophoresis, Electrophoresis, 2001, 22(4), 644–655.
118. J. Ou, T. Glawdel, C. L. Ren and J. Pawliszyn, Fabrication of a hybrid PDMS/SU-8/quartz microfluidic chip for enhancing UV absorption whole-channel imaging detection sensitivity and application for isoelectric focusing of proteins, Lab Chip, 2009, 9(13), 1926–1932.
119. M. Gilges, M. H. Kleemiss and G. Schomburg, Capillary Zone Electrophoresis Separations of Basic and Acidic Proteins Using Poly(vinyl alcohol) Coatings in Fused Silica Capillaries, Anal. Chem., 1994, 66(13), 2038–2046.
120. S. L. R. Barker, M. J. Tarlov, H. Canavan, J. J. Hickman and L. E. Locascio, Plastic Microfluidic Devices Modified with Polyelectrolyte Multilayers, Anal. Chem., 2000, 72(20), 4899–4903.
121. X. Liu, D. Erickson, D. Li and U. J. Krull, Cationic polymer coatings for design of electroosmotic flow and control of DNA adsorption, Anal. Chim. Acta, 2004, 507(1), 55–62.
122. F. Kitagawa, K. Kubota, K. Sueyoshi and K. Otsuka, One-step immobilization of cationic polymer onto a poly(methyl methacrylate) microchip for high-performance, Sci. Technol. Adv. Mater., 2006, 7(6), 558–565.
123. Y. Liu, J. C. Fanguy, J. M. Bledsoe and C. S. Henry, Dynamic coating using polyelectrolyte multilayers for chemical control of electroosmotic flow in capillary electrophoresis microchips, Anal. Chem., 2000, 72(24), 5939–5944.
124. G. Decher, J. D. Hong and J. Schmitt, Buildup of ultrathin multilayer films by a self-assembly process: III. Consecutively alternating adsorption of anionic, Thin Solid Films, 1992, 210–211, 831–835.
125. H. Katayama, Y. Ishihama and N. Asakawa, Stable Capillary Coating with Successive Multiple Ionic Polymer Layers, Anal. Chem., 1998, 70(11), 2254–2260.
126. H. Nagata, M. Tabuchi, K. Hirano and Y. Baba, Microchip electrophoretic protein separation using electroosmotic flow induced by dynamic sodium dodecyl sulfate-coating of uncoated plastic chips, Electrophoresis, 2005, 26(11), 2247–2253.
127. C. D. García, B. M. Dressen, A. Henderson and C. S. Henry, Comparison of surfactants for dynamic surface modification of poly(dimethylsiloxane) microchips, Electrophoresis, 2005, 26(3), 703–709.
128. J. S. Green and J. W. Jorgenson, Minimizing adsorption of proteins on fused silica in capillary zone electrophoresis by the addition of alkali metal salts to the buffers, Journal of Chromatography, 1989, 478, 63–70.
129. C. Gelfi, A. Vigano, M. Ripamonti, P. G. Righetti, R. Sebastiano and A. Citterio, Protein Analysis by Capillary Zone Electrophoresis Utilizing a Trifunctional Diamine for Silica Coating, Anal. Chem., 2001, 73(16), 3862–3868.
130. D. Corradini, G. Cannarsa, E. Fabbri and C. Corradini, Effects of alkylamines on electroosmotic flow and protein migration behaviour in capillary electrophoresis, J. Chromatogr., A, 1995, 709(1), 127–134.
131. P. R. Van Tassel, D. Miras, A. Hagege, M. Leroy, J.-C. Voegel and P. Schaaf, Control of Protein Adsorption in Capillary Electrophoresis via an Irreversibly Bound Protein Coating, J. Colloid Interface Sci., 1996, 183(1), 269–273.
132. N. Naruishi, Y. Tanaka, T. Higashi and S. i. Wakida, Highly efficient dynamic modification of plastic microfluidic devices using proteins in microchip capillary, J. Chromatogr., A, 2006, 1130(2), 169–174.
133. W. Schrott, Z. Slouka, P. Cervenka, J. Ston, M. Nebyla, M. Pribyl and D. Snita, Study on surface properties of PDMS microfluidic chips treated with albumin, Biomicrofluidics, 2009, 3(4), 044101–15.
134. D. G. Carlos, M. D. Brian, H. Amber and S. H. Charles, Comparison of surfactants for dynamic surface modification of poly(dimethylsiloxane) microchips, Electrophoresis, 2005, 26(3), 703–709.
135. M. F. Mora, J. Felhofer, A. Ayon and C. D. Garcia, Surfactants as a Preferred Option to Improve Separation and Electrochemical Detection in Capillary Electrophoresis, Anal. Lett., 2008, 41(2), 312–334.
136. K. K. C. Yeung and C. A. Lucy, Suppression of Electroosmotic Flow and Prevention of Wall Adsorption in Capillary Zone Electrophoresis Using Zwitterionic Surfactants, Anal. Chem., 1997, 69(17), 3435–3441.
137. Y. Luo, B. Huang, H. Wu and R. N. Zare, Controlling electroosmotic flow in poly(dimethylsiloxane) separation channels by means of prepolymer additives, Anal. Chem., 2006, 78(13), 4588–4592.
138. H. Wang, D. Wang, J. Wang, H. Wang, J. Gu, C. Han, Q. Jin, B. Xu, C. He, L. Cao, Y. Wang and J. Zhao, Application of poly(dimethylsiloxane)/glass microchip for fast electrophoretic separation of serum small, dense low-density lipoprotein, J. Chromatogr., A, 2009, 1216(35), 6343–6347.
139. S. Makino, J. A. Reynolds and C. Tanford, The Binding of Deoxycholate and Triton X-100 to Proteins, Journal of Biological Chemistry, 1973, 248(14), 4926–4932.
140. J. Z. Xing, R. Michael and E. Richards, Determination of peptidoglycan-associated protein in Escherichia coli NCIB 8545 by capillary zone electrophoresis, J. Chromatogr., A, 1996, 740(2), 273–278.
141. S. Prakash, T. M. Long, J. C. Selby, J. S. Moore and M. A. Shannon, “Click” Modification of Silica Surfaces and Glass Microfluidic Channels, Anal. Chem., 2007, 79(4), 1661–1667.
142. E. A. S. Doherty, K. D. Berglund, B. A. Buchholz, I. V. Kourkine, T. M. Przybycien, R. D. Tilton and A. E. Barron, Critical factors for high-performance physically adsorbed (dynamic) polymeric wall coatings for capillary electrophoresis of DNA, Electrophoresis, 2002, 23(16), 2766–2776.
143. A. Papra, A. Bernard, D. Juncker, N. B. Larsen, B. Michel and E. Delamarche, Microfluidic Networks Made of Poly(dimethylsiloxane), Si, and Au Coated with Polyethylene Glycol for Patterning Proteins onto Surfaces, Langmuir, 2001, 17(13), 4090–4095.
144. S. A. Soper, A. C. Henry, B. Vaidya, M. Galloway, M. Wabuyele and R. L. McCarley, Surface modification of polymer-based microfluidic devices, Anal. Chim. Acta, 2002, 470(1), 87–99.
145. S. Hu, X. Ren, M. Bachman, C. E. Sims, G. P. Li and N. L. Allbritton, Surface-Directed, Graft Polymerization within Microfluidic Channels, Anal. Chem., 2004, 76(7), 1865–1870.
146. J. Wang, A. Muck Jr., M. P. Chatrathi, G. Chen, N. Mittal, S. D. Spillman and S. Obeidat, Bulk modification of polymeric microfluidic devices, Lab Chip, 2005, 5(2), 226–230.
147. Y. Bai, C. G. Koh, M. Boreman, Y.-J. Juang, I. C. Tang, L. J. Lee and S.-T. Yang, Surface modification for enhancing antibody binding on polymer-based microfluidic device for enzyme-linked immunosorbent assay, Langmuir: The ACS Journal of Surfaces and Colloids, 2006, 22(22), 9458–9467.
148. D. Wu, B. Zhao, Z. Dai, J. Qin and B. Lin, Grafting epoxy-modified hydrophilic polymers onto poly(dimethylsiloxane) microfluidic chip to resist nonspecific protein adsorption, Lab Chip, 2006, 6(7), 942–947.
149. B. Wang, L. Chen, Z. Abdulali-Kanji, J. H. Horton and R. D. Oleschuk, Aging Effects on Oxidized and Amine-Modified Poly(dimethylsiloxane) Surfaces Studied with Chemical Force Titrations: Effects on Electroosmotic Flow Rate in Microfluidic Channels, Langmuir, 2003, 19(23), 9792–9798.
150. Y. Bai, W.-C. Huang and S.-T. Yang, Enzyme-linked immunosorbent assay of Escherichia coli O157:H7 in surface enhanced poly(methyl methacrylate) microchannels, Biotechnol. Bioeng., 2007, 98(2), 328–339.
151. J. H. Lee, B. J. Jeong and H. B. Lee, Plasma protein adsorption and platelet adhesion onto comb-like PEO gradient surfaces, J. Biomed. Mater. Res., 1997, 34(1), 105–114.
152. H. Chen, M. A. Brook and H. Sheardown, Silicone elastomers for reduced protein adsorption, Biomaterials, 2004, 25(12), 2273–2282.
153. J. Gaudioso and H. G. Craighead, Characterizing electroosmotic flow in microfluidic devices, J. Chromatogr., A, 2002, 971(1–2), 249–253.
154. T. J. Johnson, D. Ross, M. Gaitan and L. E. Locascio, Laser Modification of Preformed Polymer Microchannels: Application To Reduce Band Broadening around Turns Subject to Electrokinetic Flow, Anal. Chem., 2001, 73(15), 3656–3661.
155. X. Ren, M. Bachman, C. Sims, G. P. Li and N. Allbritton, Electroosmotic properties of microfluidic channels composed of poly(dimethylsiloxane), J. Chromatogr., B: Biomed. Sci. Appl., 2001, 762(2), 117–125.
156. K. C. Popat and T. A. Desai, Poly(ethylene glycol) interfaces: an approach for enhanced performance of microfluidic systems, Biosens. Bioelectron., 2004, 19(9), 1037–1044.
157. R. A. D'Sa and B. J. Meenan, Chemical Grafting of Poly(ethylene glycol) Methyl Ether Methacrylate onto Polymer Surfaces by Atmospheric Pressure Plasma Processing, Langmuir, 2010, 26(3), 1894–1903.
158. K. Tsougeni, D. Papageorgiou, A. Tserepi and E. Gogolides, “Smart” polymeric microfluidics fabricated by plasma processing: controlled wetting, capillary filling and hydrophobic valving, Lab Chip, 2010, 10(4), 462–469.
159. H. Yasuda, Plasma polymerization, Academic Press, 1985, p. 432.
160. P. K. Chu, J. Y. Chen, L. P. Wang and N. Huang, Plasma-surface modification of biomaterials, Mater. Sci. Eng., R, 2002, 36(5–6), 143–206.
161. L. S. McArthur and M. K. McLean, Surface Modification, in Encyclopedia of Biomaterials and Biomedical Engineering, ed. Gary E. Wnek and G. L. B., Marcel Dekker, New York, 2004, vol. 2, p. 1421.
162. H. Muguruma, Plasma-polymerized films for biosensors II, TrAC, Trends Anal. Chem., 2007, 26(5), 433–443.
163. S. Bouaidat, C. Berendsen, P. Thomsen, S. G. Petersen, A. Wolff and J. Jonsmann, Micro patterning of cell and protein non-adhesive plasma polymerized coatings for biochip applications, Lab Chip, 2004, 4(6), 632–637.
164. O. D. Krishna, K. Kim and Y. Byun, Covalently grafted phospholipid monolayer on silicone catheter surface for reduction in platelet adhesion, Biomaterials, 2005, 26(34), 7115–7123.
165. H. Thissen, G. Johnson, P. G. Hartley, P. Kingshott and H. J. Griesser, Two-dimensional patterning of thin coatings for the control of tissue outgrowth, Biomaterials, 2006, 27(1), 35–43.
166. B. Gupta, C. Plummer, I. Bisson, P. Frey and J. Hilborn, Plasma-induced graft polymerization of acrylic acid onto poly(ethylene terephthalate) films: characterization and human, Biomaterials, 2002, 23(3), 863–871.
167. G. P. Löpez, B. D. Ratner, C. D. Tidwell, C. L. Haycox, R. J. Rapoza and T. A. Horbett, Glow discharge plasma deposition of tetraethylene glycol dimethyl ether for fouling-resistant biomaterial surfaces, J. Biomed. Mater. Res., 1992, 26(4), 415–439.
168. G. P. López and B. D. Ratner, Substrate temperature effects on film chemistry in plasma depositions of organics. II. Polymerizable precursors, J. Polym. Sci., Part A: Polym. Chem., 1992, 30(11), 2415–2425.
169. G. J. Leggett, B. D. Ratner and J. C. Vickerman, Characterization of plasma-deposited styrene films by XPS and static SIMS, Surf. Interface Anal., 1995, 23(1), 22–28.
170. C. M. Chan, T. M. Ko and H. Hiraoka, Polymer surface modification by plasmas and photons, Surf. Sci. Rep., 1996, 24(1–2), 1–54.
171. M. Loughran, S. W. Tsai, K. Yokoyama and I. Karube, Simultaneous iso-electric focusing of proteins in a micro-fabricated capillary coated with hydrophobic and hydrophilic, Curr. Appl. Phys., 2003, 3(6), 495–499.
172. S.-W. Tsai, M. Loughran, A. Hiratsuka, K. Yano and I. Karube, Application of plasma-polymerized films for isoelectric focusing of proteins in a capillary electrophoresis chip, Analyst, 2003, 128(3), 237–244.
173. V. Barbier, M. Tatoulian, H. Li, F. Arefi-Khonsari, A. Ajdari and P. Tabeling, Stable modification of PDMS surface properties by plasma polymerization: application to the formation of double emulsions in microfluidic systems, Langmuir: The ACS Journal of Surfaces and Colloids, 2006, 22(12), 5230–5232.
174. M. Meincken, T. A. Berhane and P. E. Mallon, Tracking the hydrophobicity recovery of PDMS compounds using the adhesive force determined by AFM force distance, Polymer, 2005, 46(1), 203–208.
175. Z. Jinwen, E. Amanda Vera and V. Nicolas Hans, Recent developments in PDMS surface modification for microfluidic devices, Electrophoresis, 2009, 31(1), 2–16.
176. I. Wong and C.-M. Ho, Surface molecular property modifications for poly(dimethylsiloxane) (PDMS) based microfluidic devices, Microfluid. Nanofluid., 2009, 7(3), 291–306.
177. J. Zhou, H. Yan, K. Ren, W. Dai and H. Wu, Convenient Method for Modifying Poly(dimethylsiloxane) with Poly(ethylene glycol) in Microfluidics, Anal. Chem., 2009, 81(16), 6627–6632.
178. X. Yan, Y. Xiao-Dong, X. Jing-Juan and C. Hong-Yuan, Bulk modification of PDMS microchips by an amphiphilic copolymer, Electrophoresis, 2007, 28(18), 3302–3307.
179. M. Salim, G. Mishra, G. J. S. Fowler, B. O'Sullivan, P. C. Wright and S. L. McArthur, Non-fouling microfluidic chip produced by radio frequency tetraglyme plasma deposition, Lab Chip, 2007, 7(4), 523–525.
180. M. Kadowaki, H. Yoshizawa, S. Mori and M. Suzuki, Plasma CVD on the inner surface of a microchannel, Thin Solid Films, 2006, 506–507, 123–127.
181. H. Yoshiki, A. Oki, H. Ogawa and Y. Horiike, Inner wall modification of a poly(ethylene terephthalate) (PET) capillary by 13.56 MHz capacitively coupled microplasma, Thin Solid Films, 2002, 407(1–2), 156–162.
182. H. Yoshiki, Thin film coatings on capillary inner walls by microplasma, Vacuum, 2009, 84(5), 559–563.
183. J. L. Lauer, J. L. Shohet, R. M. Albrecht, C. Pratoomtong, R. Murugesan, S. Esnault, J. S. Malter, U. H. von Andrian, R. D. Bathke and S. B. Shohet, Reduced adhesion of human blood platelets to polyethylene tubing by microplasma surface modification, J. Appl. Phys., 2004, 96(8), 4539–4546.
184. H. Yoshiki, A. Oki, H. Ogawa and Y. Horiike, Generation of a capacitively coupled microplasma and its application to the inner-wall modification of a poly(ethylene terephthalate) capillary, J. Vac. Sci. Technol., A, 2002, 20(1), 24–29.
185. S. Chen, J. Zheng, L. Li and S. Jiang, Strong Resistance of Phosphorylcholine Self-Assembled Monolayers to Protein Adsorption: Insights into Nonfouling Properties of Zwitterionic Materials, J. Am. Chem. Soc., 2005, 127(41), 14473–14478.
186. S. L. McArthur, K. M. McLean, P. Kingshott, H. A. W. St John, R. C. Chatelier and H. J. Griesser, Effect of polysaccharide structure on protein adsorption, Colloids Surf., B, 2000, 17, 37–48.
187. P. Kingshott and H. J. Griesser, Surfaces that resist bioadhesion, Curr. Opin. Solid State Mater. Sci., 1999, 4(4), 403–412.
188. Z. Ademovic, B. Holst, R. Kahn, I. Jørring, T. Brevig, J. Wei, X. Hou, B. Winter-Jensen and P. Kingshott, The method of surface PEGylation influences leukocyte adhesion and activation, J. Mater. Sci.: Mater. Med., 2006, 17(3), 203–211.
189. S. B. Brueggemeier, D. Wu, S. J. Kron and S. P. Palecek, Protein–acrylamide copolymer hydrogels for array-based detection of tyrosine kinase activity from cell lysates, Biomacromolecules, 2005, 6(5), 2765–2775.
190. S. Pillai, A. Arpanaei, R. L. Meyer, V. Birkedal, L. Gram, F. Besenbacher and P. Kingshott, Preventing Protein Adsorption from a Range of Surfaces Using an Aqueous Fish Protein Extract, Biomacromolecules, 2009, 10(10), 2759–2766.
191. Y. Xu, M. Takai and K. Ishihara, Phospholipid Polymer Biointerfaces for Lab-on-a-Chip Devices, Ann. Biomed. Eng., 2010, 38(6), 1938–1953.
192. K. Jang, K. Sato, K. Mawatari, T. Konno, K. Ishihara and T. Kitamori, Surface modification by 2-methacryloyloxyethyl phosphorylcholine coupled to a photolabile linker for cell micropatterning, Biomaterials, 2009, 30(7), 1413–1420.
193. J. M. Harris, Introduction to Biotechnical and Biomedical Applications of Poly(Ethylene Glycol), in Poly(ethylene glycol) Chemistry, ed. J. M. Harris, Plenum press, 1992, p. 1.
194. N. B. Graham and M. E. McNeill, Poly(ethylene oxide): Applications in Drug Delivery, in Polymeric Materials Encyclopedia, ed. J. C. Salamone, CRC Press, New York, 1996, vol. 8, p. 6042.
195. N. Iki and E. S. Yeung, Non-bonded poly(ethylene oxide) polymer-coated column for protein separation by capillary electrophoresis, J. Chromatogr., A, 1996, 731(1–2), 273–282.
196. L. Li, S. Chen, J. Zheng, B. D. Ratner and S. Jiang, Protein Adsorption on Oligo(ethylene glycol)-Terminated Alkanethiolate Self-Assembled Monolayers: The Molecular Basis for Nonfouling Behavior, J. Phys. Chem. B, 2005, 109(7), 2934–2941.
197. P. Harder, M. Grunze, R. Dahint, G. M. Whitesides and P. E. Laibinis, Molecular Conformation in Oligo(ethylene glycol)-Terminated Self-Assembled Monolayers on Gold and Silver Surfaces Determines Their Ability To Resist Protein Adsorption, J. Phys. Chem. B, 1998, 102(2), 426–436.
198. P. Kingshott, J. Wei, D. Bagge-Ravn, N. Gadegaard and L. Gram, Covalent Attachment of Poly(ethylene glycol) to Surfaces, Critical for Reducing Bacterial Adhesion, Langmuir, 2003, 19(17), 6912–6921.
199. P. Kingshott, H. Thissen and H. J. Griesser, Effects of cloud-point grafting, chain length, and density of PEG layers on competitive adsorption of ocular proteins, Biomaterials, 2002, 23(9), 2043–2056.
200. D. Beyer, W. Knoll, H. Ringsdorf, J. H. Wang, R. B. Timmons and P. Sluka, Reduced protein adsorption on plastics via direct plasma deposition of triethylene glycol monoallyl ether, J. Biomed. Mater. Res., 1997, 36(2), 181–189.
201. L. Q. Chu, W. Knoll and R. Forch, Pulsed Plasma Polymerized Di(ethylene glycol) Monovinyl Ether Coatings for Nonfouling Surfaces, Chem. Mater., 2006, 18(20), 4840–4844.
202. Y. V. Pan, T. C. McDevitt, T. K. Kim, D. Leach-Scampavia, P. S. Stayton, D. D. Denton and B. D. Ratner, Micro-Scale Cell Patterning on Nonfouling Plasma Polymerized Tetraglyme Coatings by Protein Microcontact Printing, Plasmas Polym., 2002, 7(2), 171–183.
203. M. Shen, Y. V. Pan, M. S. Wagner, K. D. Hauch, D. G. Castner, B. D. Ratner and T. A. Horbett, Inhibition of monocyte adhesion and fibrinogen adsorption on glow discharge plasma deposited tetraethylene glycol dimethyl ether, J. Biomater. Sci., Polym. Ed., 2001, 12(9), 961–978.
204. M. Shen, M. S. Wagner, D. G. Castner, B. D. Ratner and T. A. Horbett, Multivariate Surface Analysis of Plasma-Deposited Tetraglyme for Reduction of Protein Adsorption and Monocyte Adhesion, Langmuir, 2003, 19(5), 1692–1699.
205. Y. J. Wu, R. B. Timmons, J. S. Jen and F. E. Molock, Non-fouling surfaces produced by gas phase pulsed plasma polymerization of an ultra low molecular weight ethylene oxide, Colloids Surf., B, 2000, 18(3–4), 235–248.
206. C.-G. Golander, J. N. H., Kap Lim, P. Claesson, P. Stenius and J. D. Andrade, Properties of Immobilized PEG Films and the Interaction with Proteins: Experiments and Modeling, in Poly(ethylene glycol) Chemistry, ed. J. M. Harris, Plenum Press, New York, 1992, p. 221.
207. S. I. Jeon, J. H. Lee, J. D. Andrade and P. G. De Gennes, Protein–surface interactions in the presence of polyethylene oxide: I. Simplified theory, J. Colloid Interface Sci., 1991, 142(1), 149–158.
208. Y. S. Lin, V. Hlady and C. G. Golander, The surface density gradient of grafted poly(ethylene glycol): Preparation, characterization and protein adsorption, Colloids Surf., B, 1994, 3(1–2), 49–62.
209. K. Holmberg, F. Tiberg, M. Malmsten and C. Brink, Grafting with hydrophilic polymer chains to prepare protein-resistant surfaces, Colloids Surf., A, 1997, 123–124, 297–306.
210. S. Pasche, J. Voros, H. J. Griesser, N. D. Spencer and M. Textor, Effects of Ionic Strength and Surface Charge on Protein Adsorption at PEGylated Surfaces, J. Phys. Chem. B, 2005, 109(37), 17545–17552.
211. I. Szleifer, Polymers and proteins: interactions at interfaces, Curr. Opin. Solid State Mater. Sci., 1997, 2(3), 337–344.
212. K. L. Prime and G. M. Whitesides, Self-assembled organic monolayers: model systems for studying adsorption of proteins at surfaces, Science, 1991, 252(5009), 1164–1167.
213. K. Holmberg and G. Quash, Protein Adsorption in Solid-Phase Diagnostics, in Biopolymers at Interfaces, ed. M. Malmsten, Marcel Dekker, 2003, vol. 110, p. 750.
214. S. W. Lee and P. E. Laibinis, Protein-resistant coatings for glass and metal oxide surfaces derived from oligo(ethylene glycol)-terminated alkyltrichlorosilanes, Biomaterials, 1998, 19, 1669–1675.
215. F. Bretagnol, M. Lejeune, A. Papadopoulou-Bouraoui, M. Hasiwa, H. Rauscher, G. Ceccone, P. Colpo and F. Rossi, Fouling and non-fouling surfaces produced by plasma polymerization of ethylene oxide monomer, Acta Biomater., 2006, 2(2), 165–172.
216. J. Zheng, L. Li, H.-K. Tsao, Y.-J. Sheng, S. Chen and S. Jiang, Strong Repulsive Forces between Protein and Oligo (Ethylene Glycol) Self-Assembled Monolayers: A Molecular Simulation Study, Biophys. J., 2005, 89(1), 158–166.
217. K. L. Prime and G. M. Whitesides, Adsorption of proteins onto surfaces containing end-attached oligo(ethylene oxide): a model system using self-assembled monolayers, J. Am. Chem. Soc., 1993, 115(23), 10714–10721.
218. R. S. Kane, P. Deschatelets and G. M. Whitesides, Kosmotropes Form the Basis of Protein-Resistant Surfaces, Langmuir, 2003, 19(6), 2388–2391.
219. M. Schaferling and D. Kambhampati, Protein microarray surface chemistry and coupling schemes, in Protein microarray technology, ed. D. Kamchampati, Wiley, Weinheim, 2003, pp. 11–36.
220. W.-C. Lee and K. H. Lee, Applications of affinity chromatography in proteomics, Anal. Biochem., 2004, 324(1), 1–10.
221. C. You, M. Bhagawati, A. Brecht and J. Piehler, Affinity capturing for targeting proteins into micro and nanostructures, Anal. Bioanal. Chem., 2009, 393(6–7), 1563–1570.
222. K. Jana, rcaron, enková and F. Frantiscaronek, Immobilized microfluidic enzymatic reactors, Electrophoresis, 2004, 25(21–22), 3550–3563.
223. B. Lucas, F. M. Maria, L. do L. Claudimir, A. Arturo and D. G. Carlos, Lab-on-a-Chip Biosensor for Glucose Based on a Packed Immobilized Enzyme Reactor, Electroanalysis, 2007, 19(23), 2451–2456.
224. S. Kim Shyong, Britcher, Leane, Kumar, Sunil, Griesser and J. Hans, Plasma methods for the generation of chemically reactive surfaces for biomolecule immobilization and cell colonization: A review, Wiley-VCH, Weinheim, Allemagne, 2006, pp. 392–418.
225. N. L. Anderson and N. G. Anderson, The human plasma proteome: history, character, and diagnostic prospects, Molecular & cellular proteomics: MCP, 2002, 1(11), 845–867.

---