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Research Papers

Development of Fieldable Lab-on-a-Chip Systems for Detection of a Broad Array of Targets From Toxicants to Biowarfare Agents OPEN ACCESS

[+] Author and Article Information
Aaron T. Timperman

U.S. Army Engineer Research
and Development Center,
Construction Engineering Research
Lab (ERDC-CERL),
Champaign, IL 61826

Manuscript received May 14, 2013; final manuscript received September 18, 2013; published online October 18, 2013. Assoc. Editor: Shaurya Prakash.

J. Nanotechnol. Eng. Med 4(2), 020904 (Oct 18, 2013) (8 pages) Paper No: NANO-13-1029; doi: 10.1115/1.4025539 History: Received May 14, 2013; Revised September 18, 2013

In today's world, there is an ever growing need for lightweight, portable sensor systems to detect chemical toxicants and biological toxins. The challenges encountered with such detection systems are numerous, as there are a myriad of potential targets in various sample matrices that are often present at trace-level concentrations. At ERDC-CERL, the Lab-on-a-Chip (LoaC) group is working with a number of academic and small business collaborators to develop solutions to meet these challenges. This report will focus on recent advances in three distinct areas: (1) the development of a flexible platform to allow fieldable LoaC analyses of water samples, (2) cell-, organelle-, and synthetic biology-based toxicity sensors, and (3) nanofluidic/microfluidic interface (NMI) sample enrichment devices. To transition LoaC-based sensors from the laboratory bench to the field, a portable hardware system capable of operating a wide variety of microfluidic chip-based assays has been developed. As a demonstration of the versatility of this approach assays for the separation and quantitation of anionic contaminants (i.e., perchlorate), quantitation of heavy metals (Pb and Cd), and cell-based toxicity sensors have been developed and demonstrated. Sensors harboring living cells provide a rapid means of assessing water toxicity. Cell-based sensors exploit the sensitivity of a living cell to discrete changes in its environment to report the presence of toxicants. However, this sensitivity of cells to environmental changes also hinders their usability in nonlaboratory settings. Therefore, isolating intact organelles (i.e., mitochondria) offers a nonliving alternative that preserves the sensitivity of the living cells and allows the electrochemical reporting of the presence of a contaminant. Pursuing a synthetic biology approach has also allowed the development of nonliving reporting mechanisms that utilize engineered biological pathways for novel sensing and remediation applications. To help overcome the challenges associated with the detection of target species at trace-level concentrations, NMIs are being developed for the enrichment of charged species in solution. NMI concentrators can be classified as either electroosmotic flow or electrophoresis-dominant devices. Further advances in electrophoresis-dominant concentrators will aid in the analysis of samples that contain proteins and other substances prone to surface adsorption. These recent advances illustrate how LoaC systems provide a suitable platform for development of fieldable sensors to detect a broad range of chemical/biological pollutants and threats.

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Chemical and biological detection systems have a broad range of applications, including monitoring of pollutants in the environment, assuring the safety of drinking water, and performing clinical analyses. Applications of particular interest to the military include detection of toxic compounds in water, military specific compounds in the environment, chemical and biological warfare agents, and explosives. Ideally, sample analysis can be performed in the field with lightweight, rugged, and reliable detection systems usable by personnel with minimal training. Fieldable analytical systems provide the user with a rapid response so that appropriate action can be taken to mitigate threats to the environment or to human health. Furthermore, on-site analysis reduces sample degradation or alteration and decreases costs associated with sample handling, laboratory infrastructure, and delayed response to deleterious events. Present research efforts are driven by the large gap between what is needed and what is currently available. LoaC or micrototal analysis systems (μTAS) systems possess the attributes needed to move many analytical methods to the field where rapid results are needed, sample degradation may occur during transport, for sampling when in austere environments, or when on-site sampling is simply less expensive. LoaC systems provide a platform on which the many parts of a complete sample analysis can be integrated, producing sensitive and selective sensors [1,2]. For example, sample preparation, separation, and detection have been integrated onto a single chip [3-6]. In addition, LoaC systems are inherently small, making it possible to create portable and lightweight analytical systems.

An important aim is development of sensor platforms that are broadly applicable to a number of different target molecules, analysis methods, and applications. Detection of chemical toxicants and biological toxins in a complex, nonlaboratory environment presents a difficult challenge for accurate detection due to the minimal amounts of agent necessary to inflict harm. Not only are the numbers of known toxic industrial chemicals, biological, and chemical warfare agents, and environmental contaminants staggering but not all threats are known. Therefore, detection of unknown agents also presents a significant problem for the development of sensor tools. Synthetic biology further increases the number of possible biological threats by providing a simple means to reorganize genomes and create new organisms. Therefore, a great need exists for sensors that are more universal toxicity screening tools to provide broad spectrum detection of toxic compounds. Development of these all-purpose screening tools is also valuable for environmental monitoring as a means to not only detect more traditional toxic compounds but also detect any combinatorial or synergistic effects from these compounds.

Frequently, the target molecules of interest are at trace-level concentrations and, consequently, are difficult to detect. Low concentrations, which are near or below the limits of detection (LODs), place additional burden on the analytical system. When dealing with such low concentrations, there are generally two approaches to achieving the needed signal to noise: improving the detector LODs and sample enrichment prior to the analysis. Depending on the maturity of the detection system, significant gains in the LODs may not be possible, making sample enrichment an important tool in the analysis of trace-level compounds. Therefore, sample enrichment can relieve much of the burden placed on the detection system. In addition, sample enrichment can be used to decrease the sample zone length and increase the selectivity of downstream separations. Numerous techniques for sample enrichment exist, although many techniques are difficult to integrate onto a microfluidic chip. Some of the newest and most promising methods for sample enrichment exploit the unique transport properties of ions through nanoscale channels to achieve impressive sample enrichment in microfluidic/nanofluidic systems.

To address these needs, the ERDC-CERL LoaC research program is focused on addressing these challenges by providing: (1) a standard field-deployable sensing platform that is flexible and adaptable, (2) broad spectrum biological toxicity sensors, and (3) nanofluidic sample enrichment to provide detection of compounds at trace level concentrations. In the remainder of this report, we will explore each of these research challenges in greater detail.

While advances in LoaC technology have revolutionized chemical and biological analysis in a wide variety of fields, including the analysis of environmental contaminants, these systems have largely been confined to use in the laboratory by expert users. To fully exploit the advantages conferred by the small size and weight, limited reagent consumption, and rapid analysis times of microfluidic systems, it is necessary to transition these systems forward from the laboratory to the point of sample collection. For example, real time, on-site analysis of water samples obtained after a contaminant release event could provide actionable answers to key decision makers on the ground. Accelerating this decision cycle allows rapid containment of contaminant release, limiting the extent of contamination and the cost of cleanup while reducing deleterious effects to ecosystem and human health.

Furthermore, to enable widespread adoption of microfluidics-based instrumentation by a broad user base, microfluidics-based analytical tools must be readily accessible by nonexpert users, including those with no technical background. In an ideal scenario, a minimally trained user would use portable hardware coupled with microfluidic analysis chips to conduct quantitative chemical analysis of samples with push-button operating simplicity. Results would be easily interpretable, in some cases as simple as a red light/green light response, indicating concentrations above/below a preestablished threshold, respectively.

To realize this vision, instrument developers must overcome a series of significant technical and logistical hurdles related to the transition from the laboratory bench top to more austere and less controlled environments. For example, hardware must be miniaturized and ruggedized to allow transport to sampling locations, power consumption must be minimized to reduce battery weight and increase working time, and sample handling and analysis must be automated to reduce cognitive burden on users and increase the reliability of results. These issues are exacerbated by the simultaneous transition from expert users to nonexpert users who may not be able to predict, diagnose, and mitigate potential problems.

To solve these problems and bring microfluidics-based analysis to minimally trained operators in the field, ERDC-CERL has collaborated with academic and small business partners to develop the SafePort™ portable, microfluidic chip-based water analysis system (Fig. 1). In this system, a single hardware unit is capable of operating a variety of microfluidic analysis chip designs specific to the desired analysis. This modular approach has several advantages: (1) a single hardware unit capable of running a variety of assays for different analytes and applications reduces R&D and acquisition costs, (2) a single user interface and work flow for multiple assays reduces end user cognitive burden and training requirements, and (3) a unified hardware and microfluidic chip architecture provides a pathway to transition new assays, developed anywhere, from the laboratory to the field in response to end user requirements. The SafePort™ hardware chassis makes physical fluidic connection to the top surface of the polymer microfluidic analysis chips and electrical connection to the printed circuit board base of the chips via pogo pins on the chip holder. Samples and reagents are introduced to the chip via pressure-driven flow by miniature pumping hardware housed in the chassis. High voltage power supplies in the chassis allow electrokinetic flow for on-chip sample injections and electrophoretic separations. Conductivity and electrochemical detectors allow operation of assays with a variety of detection modalities; while integration of a miniaturized fluorescence detector and lenseless camera imaging system are planned in the future. The hardware chassis interfaces to a PC via a standard USB interface. Software allows push button sample analysis with automated data analysis to provide immediate and actionable answers to the user. More sophisticated users can control assay parameters in detail to allow analysis of challenging samples, unique matrices, etc.

To demonstrate the capabilities and broad versatility of this approach, an initial library of three different analysis chips has been developed: (1) microchip capillary electrophoresis-based separation and quantitation of perchlorate, (2) electrochemical quantitation of the heavy metals lead and cadmium, and (3) a living cell-based water toxicity screening chip. The design, operation, and capabilities of each chip will be described in detail below.

Perchlorate is a persistent ground and drinking water pollutant resulting from the use of Chilean fertilizers and from the manufacture of military propellants and fuels [7]. Standard perchlorate quantitation methods (e.g., EPA 314, 331, or 332) rely on ion chromatography for separation of perchlorate from other common anions such as chloride, nitrate, and sulfate, followed by conductivity or mass spectrometric detection [8]. While these methods provide sensitive detection of perchlorate, they require expensive, resource intensive, laboratory-based instrumentation and long analysis times not amenable to rapid field sampling scenarios. The perchlorate separation and quantitation methodology used by the SafePort™ system relies on a reverse flow capillary electrophoresis-based assay using a zwitterionic sulfobetaine surfactant to separate perchlorate from more abundant anions such as chloride, nitrate, and sulfate (Fig. 2) [9]. Separation is followed by contact conductivity detection via embedded microwires. Perchlorate is quantitated via addition of an internal standard, standard addition, or construction of a calibration curve in response to end user data quality requirements. Under laboratory conditions, this assay is capable of parts per billion level limits of detection in less than 2 min. Furthermore, with small adjustments to operating parameters (e.g., separation voltage and injection time) or separation chemistry (e.g., background electrolyte and surfactant), this chip design and general separation methodology can be used for analysis of a wide variety of cations and anions from matrices ranging from groundwater to extracts from surface swabs. This capability was recently applied to use existing microfluidic chips and software to perform on-site quantitation of chlorate in groundwater monitoring wells.

Heavy metal contamination of drinking and environmental waters is a major human health concern. As with perchlorate, standard methods for quantitation of heavy metals in water rely on laboratory-based techniques such as inductively coupled plasma-mass spectrometry (ICP-MS). Electrochemical methods such as anodic stripping voltammetry (ASV) provide a portable alternative to atomic absorption or ICP-based methods while providing limits of detection below regulatory action limits. Furthermore, ASV has been implemented in several microfluidic formats, demonstrating the feasibility of the approach [10,11]. The SafePort™ heavy metal quantitation chip uses anodic stripping voltammetry with in situ bismuth plating to achieve simultaneous detection of lead and cadmium in a wide variety of matrices. In this design, a microfluidic channel is used to provide fluid transport to microfabricated graphite working, pseudo-reference, and counter electrodes. A pseudo-reference electrode was chosen to avoid the stability issues (and thus short chip lifetime) typically associated with miniaturized reference electrodes. While other fieldable stripping voltammetry-based systems for heavy metal quantitation exist, the microfluidic form factor of this assay provides advantages in terms of ease of use, sample handling, and analytical performance. Use of the standard SafePort™ interface provides the user interface, training, and cognitive burden advantages discussed above. Furthermore, the microfluidic format allows enhanced mass transport to the working electrode during sample deposition via pressure-driven flow in the microfluidic channel. Parts-per-billion limits of detection are typical for this assay, with analysis times of approximately 5 min. As with the assay above, changes to the reagent mixture and operation parameters can be used to rapidly adapt this assay to new analytes in response to user needs without the need to incur design and fabrication costs for new analysis chips.

Clearly, these assays represent only a small fraction of those that can be operated by the SafePort™ hardware. For this initial demonstration, mature and robust analytical technologies were chosen to limit technical challenges to the transition from the laboratory to the field. Our laboratory, in collaboration with academic partners, is developing a series of novel assays with the intent to transition them to SafePort™ microfluidic chips.

Ongoing research developments addressing the need for rapid, portable testing equipment have reduced the hurdles of identifying and quantifying chemicals at a field testing site or at the point of collection. However, despite the increasing ability to identify and quantify a chemical without sending samples back to an off-site laboratory, difficulty remains in assessing the environmental and human health risks associated with the contaminant identified. Developments in cell-based sensors may provide the complementary technology needed to fill these critical gaps in environmental monitoring applications.

Cell-based biosensors are composed of the biological recognition element, which is immobilized living cells (either prokaryotic or eukaryotic), interfaced with a signal transducer and the analytical device that reports the output [12]. In the integrated biosensor, the toxic effects of the contaminant on the cell generate changes to the intracellular or extracellular biochemistry that are detected and translated to either electrical or optical signals [13,14]. The high sensitivity of the biosensor stems from the ability of cells to respond to minute changes in their complex environment. Importantly, cell-based biosensors report on the bioavailable fraction of the detected contaminants as well as provide information as to the toxicity of the contaminants to the cell types present, thus providing the critical biorelevance information [15]. This actionable knowledge aids in the risk assessment as well as the selection of suitable remediation options. Unlike biorecognition-based sensors that utilize antibodies, aptamers, or other analyte-specific binding mechanisms, living cell-based biosensors can respond to emerging or little known chemical or biological threats, as well as ascertain additive and synergistic effects impossible to discern using a suite of selective sensors. Therefore, cell-based biosensors can be used as a broad spectrum screening tool by exploiting the biochemical response of the living cell to the entire complex composition of a water sample [13].

In response to this need, ERDC-CERL, in collaboration with Dr. Paul Bohn at the University of Notre Dame, is developing impedance-based cellular biosensors that can report real-time changes in cellular health as a result of toxicant assault. To build the biosensor, a confluent monolayer of cells (trout gill cells) is grown on a fibronectin-coated gold electrode array (Fig. 3). The interdigitated electrode array (IdEA) device is composed of a 5-mm square pad of 5 μm interdigitated gold microelectrode fingers. Due to the morphological and chemical changes occurring in the cells upon toxicant exposure, there is a reproducible correlation between the toxicological property and the temporal changes in the measured impedance of the reporter cell monolayer.

In addition to electrochemical means of detecting cellular responses to contaminants, an optically based reporter that utilizes fluorescence resonance energy transfer (FRET) as the signal output [16] was developed in collaboration with Dr. Yingxiao Wang at the University of Illinois at Urbana-Champaign. The cells produce an engineered FRET-based reporter targeted to cellular structures, such as focal adhesion sites; thus, it is sensitive to localized changes in reactive oxygen species (Fig. 4). Therefore, these reduction–oxidation state changes during cellular interactions with the environment can be monitored dynamically.

With the attractiveness of employing cell-based sensors as rapid, universal screening tools to detect known or unknown contaminants come the inherent drawbacks of utilizing intact living cells. The survivability of the cell population can be deleteriously affected by simple changes in temperature, pH, oxygen/carbon dioxide levels, nutrients, and sterility, thus rendering the biosensor less reliable or simply unusable. Therefore, the wide spread usage of cell-based biosensors in austere environments has been greatly hindered by the difficulties in maintaining a well-defined and healthy cell population with adequate controls.

To this end, ERDC-CERL is developing technologies that harness the power of cell-based sensors but remove the necessity to maintain intact, living cells within the biosensor device. An electrochemical sensor that reports the real-time changes in the fidelity of the electron transport chain (ETC) of isolated, intact mitochondria is being developed in collaboration with Dr. Shelley Minteer at the University of Utah. It is known that different organisms have different sensitivities to different toxicants [17]. Isolating the mitochondria from cells of different organisms retains that exquisite sensitivity but eliminates the need to use the cell as the biological recognition element. The power of using intact mitochondria immobilized onto a carbon-based electrode surface comes from the ability to tap into the ETC at either the quinone pool or the cytochrome c pool (Fig. 5). The ETC is composed of four discrete complexes that can be directly affected by certain toxicants [18]. Therefore, by using the mitochondria from a broad range of organisms, a rapid screening tool can be created to provide critical information to aid in environmental risk assessments or in the determination of the appropriate remediation methods required for a particular contaminant or class of contaminants.

The interdisciplinary field of synthetic biology combines life sciences with engineering approaches to design and build novel biological functions. Among the myriad downstream applications are production of natural and environmentally friendly biomaterials, remediation of toxic products, and sustainable energy production. As a result, another focus area of our laboratory is the production of “synthetic cells,” which are polymeric vesicles that can house functional enzymes capable of carrying out reactions applicable to contaminant remediation or energy production. These synthetic cells serve to protect their enzymatic payload from harsh external environments that would denature the enzymes or kill the organisms that natively produce the enzymes. Furthermore, enzymatic functionality from several disparate organisms can be combined in a single package to engineer a series of chemical reactions not found in nature.

As an example of this approach, we are creating synthetic cell-like vesicles capable of reducing perchlorate to chloride. Through genetic engineering, pore-forming proteins can be produced that act as gates to allow the controlled import and export of a target substrate (e.g., perchlorate) and nontoxic products (i.e., O2 and Cl) in and out of the synthetic polymer vesicle. Likewise, engineering allows the overproduction of perchlorate reducing enzymes that can be safely packaged in the interior of the synthetic polymeric vesicle, thus forming the target-selective nanoreactor (Fig. 6). Therefore, using a polymer architecture that supports and orients operational cellular machinery, biologically available “off the shelf parts” can be chosen and exploited in nonnatural combinations of cell-based functions by combining this specific machinery within a single, nonliving bioreactor to reproduce complex biological functions.

ERDC-CERL is continuing to research and develop methods to extend and enhance the usability of cell-based sensors or to maximize the potential of biological systems on synthetic platforms. Since our focus is on military applications, a robust toxicity microanalysis system, capable of functioning in harsh environments, has numerous potential uses for the soldier including rapid toxicity screening of drinking water, online monitoring of water resources, as well as monitoring at military installations to ensure environmental sustainability.

NMIs have emerged as new tools for sample enrichment of charged species in solution and have received great attention due to the high speed and concentration factors in excess of 106 reported [19-41]. The nanofluidic element can be formed by a number of methods, including nanocapillary array membranes that are also referred to as nanocapillary membranes (NCMs), planar nanochannels fabricated directly into the device substrate, and nanoporous gels. At a more basic operational level, NMI concentrators can be classified into two groups: (1) those in which analyte migration is based on electroosmotic flow (EOF) and (2) those in which analyte migration is based on electrophoresis. Therefore, EOF-dominant and electrophoresis-dominant concentrators can be readily differentiated based on the direction of analyte migration. For example, anions enriched in an NMI with negative surface charge will be driven toward the cathode in an EOF-dominant device but will migrate toward the anode in an electrophoresis-dominant device. Although, both EOF-dominant concentrators and electrophoresis-dominant concentrators have been reported, they have not been clearly differentiated in the literature. The device structure and experimental conditions determine which mode the device will operate in, and high EOF devices often have a Tee configuration while electrophoresis-dominant devices often have the nanofluidic element connecting two microchannels as shown in Fig. 7. Central to the operation of both devices is that the nanofluidic element functions as an ion permselective material. A permselective material is characterized by a transference number that is not equal to unity, meaning that the anion and cation fluxes through the permselective material are not equal. The permselectivity of a nanofluidic element is dependent on a number of parameters including the nanochannel diameter and surface charge, and the solution electrolyte concentration and pH. The permselectivity of a channel increases as κα increases, where κ is the inverse Debye length and α is the radius. As κα nears unity, double layer overlap is approached, and the potential generated by the surface charge is significant across the entire nanocapillary width. Clearly, the small radius of the nanocapillaries is critical for permselectivity and observation of concentration polarization (CP). CP is caused by the passage of ionic current through a permselective material in an electrokinetic system. In both cases, zones enriched in ions and depleted in ions form on opposite sides of nanocapillaries or nanochannels, and these zones are referred to as enriched and depleted CP zones. CP allows charged species that are smaller than the nanochannels to be retained and provides a retention mechanism that is largely charged based as opposed to a size based sieving process.

EOF-dominant concentrators have grabbed many of the highlights, due largely to the large concentration factors, in excess of 106, which have been reported [24]. In this early report and many subsequent reports, a Tee design is used that allows high EOF rates past the intersection with the nanofluidic channel. The EOF-dominant concentrators rely on high net EOF, which does not pass through the nanofluidic element in many cases [24,25,34,37,38,40,41]. These devices have many similarities with EOF pumps, with the most notable exception being the formation of the CP depleted zone in the microfluidic channel. Kim et al. fabricated an EOF-dominant concentrator using a hybrid glass/poly(dimethylsiloxane) (PDMS) device and measured high EOF rates but questioned whether they were great enough to be solely responsible for the observed increase in fluorescence intensity. The mechanism of enrichment is a based on force balancing and the nonlinear local electric-field induced by the depleted CP zone. The ions are swept toward the depleted CP zone by EOF and slow as the reverse electrophoretic force increases in response to the increasing electric field. An enriched zone forms at the point where the forward EOF and the reverse electrophoresis balance [24]. Consequently, the enriched zone is separated from the nanochannel by the depleted CP zone as shown in Fig. 7(a). Kelly et al. recommended naming this the force-balanced enriched zone to clearly differentiate it from the enriched zone, which forms in low EOF concentrators [21]. A number of investigations of the mechanisms that yield sample enrichment in the EOF-dominant concentrators have been reported [24,26]. Tallarek and co-workers presented a one-dimensional model of the enrichment process and concluded that that enrichment occurs as the forward sample delivery by EOF is balanced by the reverse electrophoretic mobility, which increases as the sample moves into the CP-depleted zone [26]. EOF of the second kind, which is nonlinear in nature and arises from the very low ion concentrations in the CP-depleted zone, is thought to be responsible for generation of the extremely high EOF rates observed. Therefore, these devices are desirable for their ability to quickly enrich sample but rely on the unusually high rate of EOF for rapid sample enrichment.

Electrophoresis-dominant concentrators usually have two microchannels that are joined directly by a nanofluidic element without a tangential field across the face of NMI. With such designs, any bulk flow, including the EOF, is greatly suppressed because it must pass through the nanofluidic element. The nanofluidic channels are great flow restrictors, as volumetric pressure driven flow is directly proportional to r4, where r is the radius. Consequently, if two microchannels are connected by a well-sealed nanofluidic element, the nanofluidic element will determine the net EOF. In contrast with high EOF concentrators, the sample is enriched in the CP-enriched zone as shown in Fig. 7(b). Both NCM and etched nanochannels have been used for the fabrication of electrophoretic concentrators [19,22,26-28,36]. Zhang and Timperman published the first report of an electrophoretic dominant concentrator [19]. The device incorporates a NCM into the fluidic design of a PDMS device. Axial resolution of the enriched and depleted zones near the NCM was not achieved, due to the geometry of the vertical microfluidic channel. In another early report, Pu et al. fabricated a planar glass device that joined two microfluidic channels with a 60-nm deep nanochannel in which they noted the enriched CP and depleted CP zones that they referred to as the ion-enrichment and ion-depletion effect [22]. Other electrophoretic-dominant concentrators incorporate gels or monoliths into the fluidic designs [26-28,36].

The appeal of electrophoretic-dominant concentrators lies largely in their compatibility with microchannel coatings that reduce EOF and flow rates that ease integration in most microfluidic systems. Unwanted nonspecific surface adsorption is a ubiquitous problem with proteomic samples, because the diverse structures of proteins make it nearly impossible to optimize solubility of all proteins within a single buffer/solvent system. It has been shown that proteins severely adsorb to charged surfaces causing sample loss and band broadening. A proteomic sample concentrator must be compatible with a coating technique that reduces protein adsorption to surfaces. Two broad categories of coating techniques are dynamic and permanent coatings. Dynamic coatings must interact with the surface through nonspecific adsorption and undergo exchange with coating monomer molecules in solution. Thus, a significant concentration of the coating monomer must be maintained in the running buffer to stabilize the coating, which is often incompatible with downstream detection and MS analysis. Permanent coatings typically alter the surface through covalent linkages. Another common method of reducing protein adsorption is to coat the microchannel or capillary surface with a positive coating and to use a low pH buffer to increase the positive charge on the proteins. However, lowering the pH reduces protein solubility [42-44], leading to clogging and sample loss. Neutral and hydrophilic polymer coatings have been shown to be the most successful permanent coatings at reducing protein adsorption but simultaneously eliminate EOF by creating a neutral surface [45-47]. Therefore, concentrators that rely on high EOF for the delivery of the sample are not preferred for the analysis of real world proteomic samples.

At ERDC-CERL, we are focused on developing broadly applicable nanofluidic concentrators that are compatible with a broad range of biological samples. Therefore, compatibility with proteins is required. To this end, we are focused on determining how device characteristics affect the enrichment process and the underlying fundamental physics of mass transport through these systems. We are combining our expertise in NMI device design and fabrication with the modeling expertise of Narayana Aluru and his group at the University of Illinois at Urbana-Champaign. In devices that couple two microfluidic channels with a NCM, we have found that small changes in device fabrication can have substantial effects on device performance. In some devices, EOF and electrophoresis are nearly balanced and small changes in the device characteristics determine whether the device is EOF or electrophoresis dominant. An example is shown in Fig. 8, which clearly shows migration in opposite directions and the separation of the enriched band from the NCM by the CP-depleted zone, although the devices have the same design differences, in fabrication have yielded different operation. An aim of our current work is to determine the device characteristics that have the greatest effect on device performance. For the targeted application, the enrichment process should take less than 5 min and achieve enrichment greater than 103. Although greater enrichment can be achieved with longer enrichment times, the longer enrichment times are not suitable for our field sensing applications.

ERDC-CERL has a multidisciplinary team focused on the goal of developing rapid, robust, sensors that provide nonexpert users with actionable results to enable an accurate risk assessment or a determination of the remediation course of action to be taken. Environmental stewardship and warfighter protection drive the advances in LoaC technology that includes the SafePort™ platform and the toolbox of user selectable chips that allow broad spectrum contaminant detection as well as universal toxicity indication. To aid the detection of compounds and organisms that are present at trace levels, NMI concentrators are being developed for rapid sample enrichment. To increase the general utility and flexibility of LoaC systems, we are developing concentrators that are compatible with proteins and other compounds prone to surface adsorption.

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Zhang, Y., and Timperman, A. T., 2003, “Integration of Nanocapillary Arrays into Microfluidic Devices for Use as Analyte Concentrators,” Analyst, 128(6), pp. 537–542. [CrossRef] [PubMed]
Miller, S. A., Kelly, K. C., and Timperman, A. T., 2008, “Ionic Current Rectification and Analyte Concentration in an Asymmetric Nanofluidic/Microfluidic Interface,” Lab Chip, 8, pp. 1729–1732. [CrossRef] [PubMed]
Kelly, K. C., Miller, S. A., and Timperman, A. T., 2009, “Investigation of Zone Migration in a Current Rectifying Nanofluidic/Microfluidic Analyte Concentrator,” Anal. Chem., 81, pp. 732–738. [CrossRef] [PubMed]
Pu, Q., Yun, J., Temkin, H., and Liu, S., 2004, “Ion-Enrichment and Ion-Depletion Effect of Nanochannel Structures,” Nano Lett., 4(6), pp. 1099–1103. [CrossRef]
Kim Sun, M., Burns Mark, A., and Hasselbrink Ernest, F., 2006, “Electrokinetic Protein Preconcentration Using a Simple Glass/Poly(Dimethylsiloxane) Microfluidic Chip,” Anal. Chem., 78(14), pp. 4779–4785. [CrossRef] [PubMed]
Wang, Y.-C., Stevens, A. L., and Han, J., 2005, “Million-Fold Preconcentration of Proteins and Peptides by Nanofluidic Filter,” Anal. Chem., 77(14), pp. 4293–4299. [CrossRef] [PubMed]
Lee, J. H., Chung, S., Kim, S. J., and Han, J., 2007, “Poly(Dimethylsiloxane)-Based Protein Preconcentration Using a Nanogap Generated by Junction Gap Breakdown,” Anal. Chem., 79(17), pp. 6868–6873. [CrossRef] [PubMed]
Dhopeshwarkar, R., Crooks, R. M., Hlushkou, D., and Tallarek, U., 2008, “Transient Effects on Microchannel Electrokinetic Filtering With an Ion-Permselective Membrane,” Anal. Chem., 80(4), pp. 1039–1048. [CrossRef] [PubMed]
Dai, J., Ito, T., Sun, L., and Crooks, R. M., 2003, “Electrokinetic Trapping and Concentration Enrichment of DNA in a Microfluidic Channel,” J. Am. Chem. Soc., 125(43), pp. 13026–13027. [CrossRef] [PubMed]
Dhopeshwarkar, R., Sun, L., and Crooks, R. M., 2005, “Electrokinetic Concentration Enrichment Within a Microfluidic Device Using a Hydrogel Microplug,” Lab Chip, 5(10), pp. 1148–1154. [CrossRef] [PubMed]
Khandurina, J., Jacobson, S. C., Waters, L. C., Foote, R. S., and Ramsey, J. M., 1999, “Microfabricated Porous Membrane Structure for Sample Concentration and Electrophoretic Analysis,” Anal. Chem., 71(9), pp. 1815–1819. [CrossRef] [PubMed]
Foote, R. S., Khandurina, J., Jacobson, S. C., and Ramsey, J. M., 2005, “Preconcentration of Proteins on Microfluidic Devices Using Porous Silica Membranes,” Anal. Chem., 77(1), pp. 57–63. [CrossRef] [PubMed]
Hlushkou, D., Dhopeshwarkar, R., Crooks Richard, M., and Tallarek, U., 2008, “The Influence of Membrane Ion-Permselectivity on Electrokinetic Concentration Enrichment in Membrane-Based Preconcentration Units,” Lab Chip, 8(7), pp. 1153–1162. [CrossRef] [PubMed]
Huang, K.-D., and Yang, R.-J., 2008, “Formation of Ionic Depletion/Enrichment Zones in a Hybrid Micro-/Nano-Channel,” Microfluid. Nanofluid., 5(5), pp. 631–638. [CrossRef]
Yu, H., Lu, Y., Zhou, Y.-G., Wang, F.-B., He, F.-Y., and Xia, X.-H., 2008, “A Simple, Disposable Microfluidic Device for Rapid Protein Concentration and Purification via Direct-Printing,” Lab Chip, 8(9), pp. 1496–1501. [CrossRef] [PubMed]
Wang, Y.-C., and Han, J., 2008, “Pre-Binding Dynamic Range and Sensitivity Enhancement for Immuno-Sensors Using Nanofluidic Preconcentrator,” Lab Chip, 8(3), pp. 392–394. [CrossRef] [PubMed]
Stein, D., Deurvorst, Z., Van Der Heyden, F. H. J., Koopmans, W. J. A., Gabel, A., and Dekker, C., 2010, “Electrokinetic Concentration of DNA Polymers in Nanofluidic Channels,” Nano Lett., 10(3), pp. 765–772. [CrossRef] [PubMed]
Yamamoto, S., Hirakawa, S., and Suzuki, S., 2008, “In Situ Fabrication of Ionic Polyacrylamide-Based Preconcentrator on a Simple Poly(Methyl Methacrylate) Microfluidic Chip for Capillary Electrophoresis of Anionic Compounds,” Anal. Chem, 80(21), pp. 8224–8230. [CrossRef] [PubMed]
Lee, J. H., Song, Y.-A., and Han, J., 2008, “Multiplexed Proteomic Sample Preconcentration Device Using Surface-Patterned Ion-Selective Membrane,” Lab Chip, 8(4), pp. 596–601. [CrossRef] [PubMed]
Hoeman, K. W., Lange, J. J., Roman, G. T., Higgins, D. A., and Culbertson, C. T., 2009, “Electrokinetic Trapping Using Titania Nanoporous Membranes Fabricated Using Sol-Gel Chemistry on Microfluidic Devices,” Electrophoresis, 30(18), pp. 3160–3167. [CrossRef] [PubMed]
Zhou, K., Kovarik, M. L., and Jacobson, S. C., 2008, “Surface-Charge Induced Ion Depletion and Sample Stacking Near Single Nanopores in Microfluidic Devices,” J. Am. Chem. Soc., 130(27), pp. 8614–8616. [CrossRef] [PubMed]
Kovarik, M. L., and Jacobson, S. C., 2008, “Integrated Nanopore/Microchannel Devices for AC Electrokinetic Trapping of Particles,” Anal. Chem, 80(3), pp. 657–664. [CrossRef] [PubMed]
Kim, S. J., and Han, J., 2008, “Self-Sealed Vertical Polymeric Nanoporous-Junctions for High-Throughput Nanofluidic Applications,” Anal. Chem, 80(9), pp. 3507–3511. [CrossRef] [PubMed]
Moini, M., and Huang, H., 2004, “Application of Capillary Electrophoresis/Electrospray Ionization-Mass Spectrometry to Subcellular Proteomics of Escherichia Coli Ribosomal Proteins,” Electrophoresis, 25(13), pp. 1981–1987. [CrossRef] [PubMed]
Simpson, D. C., and Smith, R. D., 2005, “Combining Capillary Electrophoresis With Mass Spectrometry for Applications in Proteomics,” Electrophoresis, 26(7–8), pp. 1291–1305. [CrossRef] [PubMed]
Schiffer, E., Mischak, H., and Novak, J., 2006, “High Resolution Proteome/Peptidome Analysis of Body Fluids by Capillary Electrophoresis Coupled With Ms,” Proteomics, 6(20), pp. 5615–5627. [CrossRef] [PubMed]
Ostuni, E., Chapman, R. G., Liang, M. N., Meluleni, G., Pier, G., Ingber, D. E., and Whitesides, G. M., 2001, “Self-Assembled Monolayers That Resist the Adsorption of Proteins and the Adhesion of Bacterial and Mammalian Cells,” Langmuir, 17(20), pp. 6336–6343. [CrossRef]
Razunguzwa, T. T., Warrier, M., and Timperman, A. T., 2006, “ESI-MS Compatible Permanent Coating of Glass Surfaces Using Poly(Ethylene Glycol)-Terminated Alkoxysilanes for Capillary Zone Electrophoretic Protein Separations,” Anal. Chem., 78(13), pp. 4326–4333. [CrossRef] [PubMed]
Sun, X., Liu, J., and Lee, M. L., 2008, “Surface Modification of Polymer Microfluidic Devices Using in-Channel Atom Transfer Radical Polymerization,” Electrophoresis, 29(13), pp. 2760–2767. [CrossRef] [PubMed]
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References

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Chinowsky, T. M., Grow, M. S., Johnston, K. S., Nelson, K., Edwards, T., Fu, E., and Yager, P., 2007, “Compact, High Performance Surface Plasmon Resonance Imaging System,” Biosens. Bioelectron., 22(9–10), pp. 2208–2215. [CrossRef] [PubMed]
U.S.E.P. Agency, 2011, “Final Regulatory Determination for Perchlorate, 5/6,” https://www.federalregister.gov/articles/2011/02/11/2011-2603/drinking-water-regulatory-determination-on-perchlorate
Urbansky, E. T., 2000, “Quantitation of Perchlorate Ion: Practices and Advances Applied to the Analysis of Common Matrices,” Crit. Rev. Anal. Chem., 30(4), pp. 311–343. [CrossRef]
Gertsch, J. C., Noblitt, S. D., Cropek, D. M., and Henry, C. S., 2010, “Rapid Analysis of Perchlorate in Drinking Water at Parts Per Billion Levels Using Microchip Electrophoresis,” Anal. Chem., 82(9), pp. 3426–3429. [CrossRef] [PubMed]
Zou, Z. W., Jang, A., Macknight, E., Wu, P. M., Do, J., Bishop, P. L., and Ahn, C. H., 2008, “Environmentally Friendly Disposable Sensors With Microfabricated on-Chip Planar Bismuth Electrode for In Situ Heavy Metal Ions Measurement,” Sens. Actuators B, 134(1), pp. 18–24. [CrossRef]
Wang, J., Polsky, R., Tian, B. M., and Chatrathi, M. P., 2000, “Voltammetry on Microfluidic Chip Platforms,” Anal. Chem., 72(21), pp. 5285–5289. [CrossRef] [PubMed]
Rogers, K. R., 2006, “Recent Advances in Biosensor Techniques for Environmental Monitoring,” Anal. Chim. Acta, 568(1–2), pp. 222–231. [CrossRef] [PubMed]
Banerjee, P., Franz, B., and Bhunia, A. K., 2010, “Mammalian Cell-Based Sensor System,” Adv. Biochem. Eng. Biotechnol., 117, pp. 21–55. [CrossRef] [PubMed]
Wang, P., 2010, Cell-Based Biosensors Principles and Applications, Artech House, Norwood, MA.
Lagarde, F., and Jaffrezic-Renault, N., 2011, “Cell-Based Electrochemical Biosensors for Water Quality Assessment,” Anal. Bioanal. Chem., 400(4), pp. 947–964. [CrossRef] [PubMed]
Lin, L. J., Grimme, J. M., Sun, J., Lu, S., Gai, L., Cropek, D. M., and Wang, Y., 2013, “The Antagonistic Roles of Pdgf and Integrin Alphavbeta3 in Regulating Ros Production at Focal Adhesions,” Biomaterials, 34(15), pp. 3807–3815. [CrossRef] [PubMed]
Vicente, J. A., Peixoto, F., Lopes, M. L., and Madeira, V. M., 2001, “Differential Sensitivities of Plant and Animal Mitochondria to the Herbicide Paraquat,” J. Biochem. Mol. Toxicol., 15(6), pp. 322–330. [CrossRef] [PubMed]
Fernandes, M. A., Santos, M. S., Alpoim, M. C., Madeira, V. M., and Vicente, J. A., 2002, “Chromium(Vi) Interaction With Plant and Animal Mitochondrial Bioenergetics: A Comparative Study,” J. Biochem. Mol. Toxicol., 16(2), pp. 53–63. [CrossRef] [PubMed]
Zhang, Y., and Timperman, A. T., 2003, “Integration of Nanocapillary Arrays into Microfluidic Devices for Use as Analyte Concentrators,” Analyst, 128(6), pp. 537–542. [CrossRef] [PubMed]
Miller, S. A., Kelly, K. C., and Timperman, A. T., 2008, “Ionic Current Rectification and Analyte Concentration in an Asymmetric Nanofluidic/Microfluidic Interface,” Lab Chip, 8, pp. 1729–1732. [CrossRef] [PubMed]
Kelly, K. C., Miller, S. A., and Timperman, A. T., 2009, “Investigation of Zone Migration in a Current Rectifying Nanofluidic/Microfluidic Analyte Concentrator,” Anal. Chem., 81, pp. 732–738. [CrossRef] [PubMed]
Pu, Q., Yun, J., Temkin, H., and Liu, S., 2004, “Ion-Enrichment and Ion-Depletion Effect of Nanochannel Structures,” Nano Lett., 4(6), pp. 1099–1103. [CrossRef]
Kim Sun, M., Burns Mark, A., and Hasselbrink Ernest, F., 2006, “Electrokinetic Protein Preconcentration Using a Simple Glass/Poly(Dimethylsiloxane) Microfluidic Chip,” Anal. Chem., 78(14), pp. 4779–4785. [CrossRef] [PubMed]
Wang, Y.-C., Stevens, A. L., and Han, J., 2005, “Million-Fold Preconcentration of Proteins and Peptides by Nanofluidic Filter,” Anal. Chem., 77(14), pp. 4293–4299. [CrossRef] [PubMed]
Lee, J. H., Chung, S., Kim, S. J., and Han, J., 2007, “Poly(Dimethylsiloxane)-Based Protein Preconcentration Using a Nanogap Generated by Junction Gap Breakdown,” Anal. Chem., 79(17), pp. 6868–6873. [CrossRef] [PubMed]
Dhopeshwarkar, R., Crooks, R. M., Hlushkou, D., and Tallarek, U., 2008, “Transient Effects on Microchannel Electrokinetic Filtering With an Ion-Permselective Membrane,” Anal. Chem., 80(4), pp. 1039–1048. [CrossRef] [PubMed]
Dai, J., Ito, T., Sun, L., and Crooks, R. M., 2003, “Electrokinetic Trapping and Concentration Enrichment of DNA in a Microfluidic Channel,” J. Am. Chem. Soc., 125(43), pp. 13026–13027. [CrossRef] [PubMed]
Dhopeshwarkar, R., Sun, L., and Crooks, R. M., 2005, “Electrokinetic Concentration Enrichment Within a Microfluidic Device Using a Hydrogel Microplug,” Lab Chip, 5(10), pp. 1148–1154. [CrossRef] [PubMed]
Khandurina, J., Jacobson, S. C., Waters, L. C., Foote, R. S., and Ramsey, J. M., 1999, “Microfabricated Porous Membrane Structure for Sample Concentration and Electrophoretic Analysis,” Anal. Chem., 71(9), pp. 1815–1819. [CrossRef] [PubMed]
Foote, R. S., Khandurina, J., Jacobson, S. C., and Ramsey, J. M., 2005, “Preconcentration of Proteins on Microfluidic Devices Using Porous Silica Membranes,” Anal. Chem., 77(1), pp. 57–63. [CrossRef] [PubMed]
Hlushkou, D., Dhopeshwarkar, R., Crooks Richard, M., and Tallarek, U., 2008, “The Influence of Membrane Ion-Permselectivity on Electrokinetic Concentration Enrichment in Membrane-Based Preconcentration Units,” Lab Chip, 8(7), pp. 1153–1162. [CrossRef] [PubMed]
Huang, K.-D., and Yang, R.-J., 2008, “Formation of Ionic Depletion/Enrichment Zones in a Hybrid Micro-/Nano-Channel,” Microfluid. Nanofluid., 5(5), pp. 631–638. [CrossRef]
Yu, H., Lu, Y., Zhou, Y.-G., Wang, F.-B., He, F.-Y., and Xia, X.-H., 2008, “A Simple, Disposable Microfluidic Device for Rapid Protein Concentration and Purification via Direct-Printing,” Lab Chip, 8(9), pp. 1496–1501. [CrossRef] [PubMed]
Wang, Y.-C., and Han, J., 2008, “Pre-Binding Dynamic Range and Sensitivity Enhancement for Immuno-Sensors Using Nanofluidic Preconcentrator,” Lab Chip, 8(3), pp. 392–394. [CrossRef] [PubMed]
Stein, D., Deurvorst, Z., Van Der Heyden, F. H. J., Koopmans, W. J. A., Gabel, A., and Dekker, C., 2010, “Electrokinetic Concentration of DNA Polymers in Nanofluidic Channels,” Nano Lett., 10(3), pp. 765–772. [CrossRef] [PubMed]
Yamamoto, S., Hirakawa, S., and Suzuki, S., 2008, “In Situ Fabrication of Ionic Polyacrylamide-Based Preconcentrator on a Simple Poly(Methyl Methacrylate) Microfluidic Chip for Capillary Electrophoresis of Anionic Compounds,” Anal. Chem, 80(21), pp. 8224–8230. [CrossRef] [PubMed]
Lee, J. H., Song, Y.-A., and Han, J., 2008, “Multiplexed Proteomic Sample Preconcentration Device Using Surface-Patterned Ion-Selective Membrane,” Lab Chip, 8(4), pp. 596–601. [CrossRef] [PubMed]
Hoeman, K. W., Lange, J. J., Roman, G. T., Higgins, D. A., and Culbertson, C. T., 2009, “Electrokinetic Trapping Using Titania Nanoporous Membranes Fabricated Using Sol-Gel Chemistry on Microfluidic Devices,” Electrophoresis, 30(18), pp. 3160–3167. [CrossRef] [PubMed]
Zhou, K., Kovarik, M. L., and Jacobson, S. C., 2008, “Surface-Charge Induced Ion Depletion and Sample Stacking Near Single Nanopores in Microfluidic Devices,” J. Am. Chem. Soc., 130(27), pp. 8614–8616. [CrossRef] [PubMed]
Kovarik, M. L., and Jacobson, S. C., 2008, “Integrated Nanopore/Microchannel Devices for AC Electrokinetic Trapping of Particles,” Anal. Chem, 80(3), pp. 657–664. [CrossRef] [PubMed]
Kim, S. J., and Han, J., 2008, “Self-Sealed Vertical Polymeric Nanoporous-Junctions for High-Throughput Nanofluidic Applications,” Anal. Chem, 80(9), pp. 3507–3511. [CrossRef] [PubMed]
Moini, M., and Huang, H., 2004, “Application of Capillary Electrophoresis/Electrospray Ionization-Mass Spectrometry to Subcellular Proteomics of Escherichia Coli Ribosomal Proteins,” Electrophoresis, 25(13), pp. 1981–1987. [CrossRef] [PubMed]
Simpson, D. C., and Smith, R. D., 2005, “Combining Capillary Electrophoresis With Mass Spectrometry for Applications in Proteomics,” Electrophoresis, 26(7–8), pp. 1291–1305. [CrossRef] [PubMed]
Schiffer, E., Mischak, H., and Novak, J., 2006, “High Resolution Proteome/Peptidome Analysis of Body Fluids by Capillary Electrophoresis Coupled With Ms,” Proteomics, 6(20), pp. 5615–5627. [CrossRef] [PubMed]
Ostuni, E., Chapman, R. G., Liang, M. N., Meluleni, G., Pier, G., Ingber, D. E., and Whitesides, G. M., 2001, “Self-Assembled Monolayers That Resist the Adsorption of Proteins and the Adhesion of Bacterial and Mammalian Cells,” Langmuir, 17(20), pp. 6336–6343. [CrossRef]
Razunguzwa, T. T., Warrier, M., and Timperman, A. T., 2006, “ESI-MS Compatible Permanent Coating of Glass Surfaces Using Poly(Ethylene Glycol)-Terminated Alkoxysilanes for Capillary Zone Electrophoretic Protein Separations,” Anal. Chem., 78(13), pp. 4326–4333. [CrossRef] [PubMed]
Sun, X., Liu, J., and Lee, M. L., 2008, “Surface Modification of Polymer Microfluidic Devices Using in-Channel Atom Transfer Radical Polymerization,” Electrophoresis, 29(13), pp. 2760–2767. [CrossRef] [PubMed]

Figures

Grahic Jump Location
Fig. 1

(Top) Early prototype of SafePort™ hardware showing electronic component housing, chip carrier, and microfluidic chip. (Bottom) SafePort™ compatible microfluidic chip composed of hot embossed polymer on a printed circuit board base.

Grahic Jump Location
Fig. 2

Electropherograms showing separation of drinking water samples spiked with 0.12 ppm PDS and concentrations of perchlorate between 1 and 1000 ppb. Conditions: −350 V/cm, 10 s injection, background electrolyte = 10 mM nicotinic acid, 1.0 mM TDAPS, pH 3.6. Reprinted with permission from Gertsch et al. [9]. Copyright (2010) American Chemical Society.

Grahic Jump Location
Fig. 3

Schematic representation of the IdEA for impedance-based detection of contaminants in water. (Left) Representation of the working sensor device showing positions of all gold electrodes, the IdEA pad where the monolayer of cells are seeded onto the extracellular matrix-coated gold electrode array. (Middle) Monolayer of trout gill cells grown on the IdEA pad. (Right) Live (light) versus dead (dark) cell staining of the cells following toxicant exposure.

Grahic Jump Location
Fig. 4

Individual mouse embryonic fibroblast cells harboring FRET-based reporter targeted to focal adhesion sites. (Left) The ECFP (donor)/YPet (acceptor) ratio images in response to the oxidizing agent, diamide, over time. (Right) Time course of the normalized ECFP/YPet ratio upon treatment with diamide (0.5 mM).

Grahic Jump Location
Fig. 5

Schematic representation of a mitochondrial bioelectrode showing electron and ATP production during substrate (pyruvate or fatty acid) oxidation through the four complexes of the electron transport chain and ATPase

Grahic Jump Location
Fig. 6

Synthetic polymer nanoreactor. A polymer-based shell with protein gates for controlled influx and efflux. The encapsulated proteins are responsible for transformation of the targeted substrate, perchlorate. Image created by Dr. Manish Kumar, Pennsylvania State University.

Grahic Jump Location
Fig. 7

Examples of high (a) and low EOF (b) NMI sample concentrators are shown. The signs in the reservoirs near the microchannel ends show the polarity of the applied electric field. In both cases, the permselectivity of the nanofluidic element and electric field drive the formation of the depleted and enriched CP zones on opposite sides of the nanofluidic element. In the high EOF system, the microchannel EOF is greater in magnitude than the electrophoretic mobility of the anions, driving the anions to the interface of the bulk solution with the depleted CP zone. At this interface, the reverse electrophoretic velocity of the analyte increases as the local electric field increases, and the analyte is enriched in a forced balanced zone. In the low EOF system, the nanofluidic element is in the flow path and sufficiently reduces the EOF, and electrophoresis drives the analyte toward the nanofluidic element where it is concentrated in the enriched CP zone.

Grahic Jump Location
Fig. 8

Two similar NMI concentrators with integrated NCMs. (a) The anionic fluorescein is transported towards the negative electrode by EOF. The separation between the NCM and the enriched zone is NCM is caused by presence of the CP depleted zone. (b) The EOF is lower and the fluorescein migrates toward the NCM by electrophoresis. The highest intensity is in the vertical microchannel via.

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