Jump to ContentJump to Main Navigation
Show Summary Details
The one-stop-shop for nano science.

nano Online

Physics, Chemistry and Materials Science at the Nanoscale

More options …


Search publication

Open Access

1 Overview of point-of-care technologies

Patient care and diagnosis revolves around understanding an individual person’s unique and complex physiology and is historically done through the collection and analysis of bodily fluids. Blood, urine, and sweat (BUS) are the most common biological samples collected from patients and used in laboratory testing. However, various other bodily samples, such as cerebral spinal fluid or sputum, can also be used for testing but can be much more difficult to obtain from the patient or are less well studied in terms of their correlation with the known clinical blood biomarkers [1].

Results obtained from BUS assays provide physicians with valuable insights for improved diagnoses, monitoring, and treating of patients [2]. Therefore, it is understandable that over 13 billion in vitro diagnostic tests (IVDs) are performed annually [3]. To drive healthcare forward and facilitate more predictive, personalized medical care, the testing of biological fluid using point-of-care (POC) technologies will become increasingly paramount [4], [5].

In the US, 86% of all IVDs are performed off-site at independent laboratories or centralized hospitals [3]. Patients are typically referred to a separate location where biological samples can be collected and analyzed by a phlebotomist and trained technician, respectively. Laboratory testing of patients in underserved areas including developing countries presents even greater challenges due to the limited access to laboratory equipment, clean water, and consistent electricity. Patient samples are typically sent to off-site locations capable of performing such tests, which runs the risk of samples being contaminated, lost, or mislabeled [6]. Many patients in underserved settings are unable to return to city clinics for multiple appointments and often never receive results, only the initial medication, further prolonging proper diagnosis and treatment [7].

The hassle of off-site laboratory testing and the stress of waiting for results motivate the need and demonstrate opportunity for improvement and innovation in this area of healthcare. Recent developments in POC technology focus on creating rapid diagnostic tests (RDTs) where results are obtained minutes after collection, yet many of these tests are not as accurate as their laboratory-based equivalents. Enhancing these tests through the development of POC technologies that involve automated sample preparation, such as microfluidic lab-on-a-chip (LOC) devices have the potential to remove the need for complex laboratories and experienced personnel. If they can be made equally as accurate, robust, and easy to use this could improve overall health quality by bringing BUS test results to the patient’s bedside, into the home, inside the ambulance, or on the field [8].

2 Defining the ideal characteristics of a POC technology

A POC technology should ideally be accurate, portable, simple to use [2], and easy to read a positive or negative result; require little or no sample preparation; provide timely results; and be cost-effective [9]. The goal of POC technology is to provide a user-friendly interface, without the need of expert technician support or complex analysis, while meeting the current clinical chemistry standards for IVDs as set by the AACC [10], [11].

The World Health Organization (WHO) has defined a set of parameters that outline the specifications for POC technology in developing areas. This outline was originally created for the development of portable devices for human immunodeficiency virus (HIV) testing. The parameters are commonly referred to as the ASSURED criteria [12]. According to the WHO, POC technology must be affordable for patients without creating financial strain, sensitive to detect the biological target molecules at physiologic concentrations comparable to the central laboratory result, specific to only the target biomolecule and impervious to comparable molecules, user-friendly and easy to use with little or no training or expertise necessary, rapid and robust to provide timely results using samples that can be handled under the POC condition needed for the setting, equipment that is large and inconvenient should be avoided, and deliverable to patients in need at their location. These parameters have been well accepted and are the general standard criteria for classification as a POC technology [13].

2.1 Current commercial POC technologies

There are POC devices that are commercially available. The most familiar example is the glucose meter for managing diabetes, a chronic illness effecting 29 million people in the US alone [14], [15]. It is defined as a group of metabolic diseases where ultimately the body’s pancreas does not produce enough insulin or does not properly respond to insulin produced, resulting in high blood sugar levels over a prolonged period. Glucose meters and other POC devices utilize an assortment of methods for detecting and monitoring biomarkers including electrochemical [16], [17], [18], [19], [20], magnetic [21], [22], [23], [24], [25], [26], [27], [28], [29], [30], optical [31], [32], [33], [34], label-free spectroscopic analysis [35], [36], [37], [38], [39], [40], [41], [42], [43], colorimetric [44], [45], [46], [47], [48], [49], and plasmonic nanoparticle based sensors [50], [51], [52]. Generally, electrochemical detection uses potentiometric, amperometric, and impedimetric measurements in conjunction with electroactive tags or free flowing electroactive analytes [17], [18], [19], [20]. Many of the commercial glucose detection monitors One Touch Ultra (Johnson & Johnson, Wayne, PA, USA), Arkray (Edina, MN, USA), Ascensia Contour (Ascensia, Parsippany, NJ, USA), BD Test Strip (Becton Dickinson, Franklin Lakes, NJ, USA), FreeStyle and Precision Xtra (Abbott Laboratories, Chicago, IL, USA), TrueTrack Smart System (Trividia, Fort Lauderdale, FL, USA), Accu-chek Aviva (Roche, Basel, Switzerland), etc. [15], [53], [54] are examples of electrochemical and colorimetric devices that use test strips. In addition, the i-STAT monitor (Abbott Laboratories, Chicago, IL, USA) is an electrochemical approach that uses individual microfluidic cartridges for monitoring lactate, pH, pO2, pCO2, electrolytes, and hematology [55].

A newer commercial POC approach that uses magnetic nanoparticles, an antibody immunoassay, and optical detection to measure troponin, one biomarker for detection of myocardial infarction (heart attack), was recently developed by Philips (Handheld Minicare I-20). The system was created for use in emergency departments and provides results in 10 min from a droplet of blood instead of the typical 1-h wait incurred using central laboratories [56], [57].

Other examples include the mChip, a device that tests whole blood for HIV using a combined microfluidic and protein-based immunoassay approach [58], and the Ativa Micro Lab (Ativa Medical, St. Paul, MN, USA), which was recently shown to execute a number of distinct blood tests with the use of a single device [59], [60], [61]. One upcoming technology is Ativa’s portable Micro Lab, which has the potential to perform up to 25 distinct blood tests utilizing its various disposable test cards for each test. The Micro Lab system uses microfluidics, flow cytometry, electrochemistry, and colorimetric readouts in combination with established imaging techniques to conduct and relay important diagnostic information. While not yet approved by the FDA or commercially available and more of a bench-top reader rather than a hand-held POC device, it shows promise as a potential portable and user-friendly instrument based on a microfluidic cartridge suited to measure several tests.

One of the most popular emerging POC techniques that are commercially available for some applications are based on lateral flow assays (LFAs), which use colorimetric barcodes that can be viewed with the naked eye, absorbance measurements with a cell phone, or fluorescence techniques to quantify various biomarkers. The most popular LFA is the pregnancy test First Response (Church & Dwight Co., Inc., Trenton, NJ, USA), Clearblue (Swiss Precision Diagnostics GmbH, Geneva Switzerland) etc. [1], [4], [62]. The technology is based on porous paper that contains an immunoassay, gold nanoparticles, and a porous membrane capable of transporting fluid (e.g. urine). As the solution migrates down the paper, the target molecule, sometimes referred to as the antigen, chemically reacts with its antibody that has been immobilized on a particle surface. The particles can then be trapped downstream in the strip with another antibody and read with an optical reader for quantification or a yes/no readout can be obtained by visual inspection [63].

Although there are many POC devices commercially available, they are primarily designed to produce semi-quantitative or quantitative information for a single analyte per test [10]. Thus, the common limitations of commercially available POC devices are accuracy, variations of cost per test, and the inability to simultaneously monitor multiple analytes. All of these factors are important and often required to make definitive diagnoses [61], [64], [65], [66], [67]. As a consequence, research and engineering efforts are still required to address these factors to improve POC technology.

2.2 Optical analyses for facilitating POC technologies

Current research efforts are directed at developing POC technologies with the necessary performance to be useful for diagnostics. In addition, physicians and clinicians are interested in the emergence of POC devices capable of simultaneously sensing multiple biomarkers and analytes characteristic of specific disease states [9], [68]. The additional push for one technology to monitor several biomarkers simultaneously for more effective and universal disease diagnosis is motivating recent advancements in multiplexed sensing capabilities.

Many optical approaches have been utilized for POC applications including fluorescence [69], [70], [71], [72], luminescence, absorbance, Forster energy transfer (FRET) [73], bioluminescence energy transfer (BRET) [74], [75], [76], surface-plasmon resonance (SPR) [77], [78], resonant Rayleigh scattering [79], [80], [81], [82], geometric scattering [1], and Raman spectroscopic approaches [83], [84], [85], [86], [87], [88], [89]. Advancements in biochemical sensing methods, nanotechnology, and the miniaturization of optics have been key in improving spectroscopic platforms for biomarker detection.

In many cases, optical approaches are more successful in simultaneously sensing multiple analytes through the use of photoactive dyes conjugated to biorecognition ligands [33]. Indirect sensing through the identification of optically active molecules, including chromophores and fluorophores or other reporter molecules, has improved the sensitivity and multiplexing capabilities of techniques like Raman or fluorescence spectroscopy while simultaneously lowering the limits of detection (LODs) and broadening the dynamic range of spectroscopic analysis [31], [90]. This has led to fluorescence-based POC devices, in particular, as being one of the most widely explored and currently utilized optical modalities for the detection of multiple analytes on a single test. Examples of fluorescence-based POC devices being explored include LOC microfluidic immunoassays and early disease and cancer detection through biomarker isolation [91], [92]. In terms of the equipment and optics, in recent years, there has been a significant advancement in portable handheld fluorescence spectrometers. Various commercial groups have successfully compacted bench-top fluorescence systems into portable and easy-to-use hand-held devices. Cutting edge advancements in affordable optical sensing and imaging have also been achieved, most notably with the development of cellphone-based designs [58], [93] that can have major healthcare impact in low resources settings including the developing world.

Fluorescence POC devices do have some limitations, particularly relative to their multiplexing capabilities due to the dyes’ broad emission spectrums, which can overlap when using multiple dyes. Furthermore, the need for different excitation wavelengths to excite multiple dyes can pose additional challenges [94], [95].

The use of dyes or reporter molecules has also become popular in other spectroscopic platforms beyond fluorescence. Most interesting has been the use of multiple fluorescent dyes, photosensitizers, or other reporter molecules with Raman spectroscopy. The spectra of the same dye reporter obtained using Raman produce spectral peaks with a full width half maximum 10–100 times narrower than the peaks typically observed using fluorescence [27]. These thin spectral bands provide the capacity for SERS to be used for multiplexed detection of several biomarkers from a single measurement [96].

3 Surface enhanced Raman spectroscopy (SERS) and its potential benefits

Raman spectroscopy provides a “chemical fingerprint” as it relies on loss (Stokes) or gain (anti-Stokes) in energy of an inelastically scattered photon due to a molecular vibrational event. Traditional spontaneous Raman scattering is intrinsically inefficient compared to elastic Rayleigh scattering, as it relies on the polarizability, or Raman cross-section, of the analyte. Even when probing analytes such as MBA, DTNB, and BPE, which have aromatic rings (containing benzene) whose pi-bonds can be easily delocalized around the ring, producing distinct vibrational modes [97], the probability of a photon to be scattered inelastically is only ~1 in 108 [98].

SERS is one technique that can enhance the Raman signal by several orders of magnitude by amplifying the electron cloud density around metallic nanostructures [99], [100], [101]. SERS signal enhancement is achieved through two distinct mechanisms: electromagnetic enhancement and chemical enhancement [100], [102], [103], [104]. For both mechanisms to occur simultaneously, an analyte must be adsorbed onto or reside very close to a dielectric (often metallic) surface [90]. Electromagnetic enhancement utilizes metallic nanostructures with local surface plasmon resonance (LSPR) wavelengths in resonance with the Raman excitation source. When this coupling occurs, plasmons (oscillating conduction band electrons) increase the local electron density, thus improving the likelihood of inelastic scatter events to occur when the analyte is near the particle surface [105], [106]. For chemical enhancement to occur, a molecule must be able to be bound directly to the surface of the metal so that there is a charge-transfer structure generated, i.e. an electron-hole pair that can mediate the transfer of energy from the metal directly to the molecular bonds of the analyte [103], [104].

Typical SERS signal enhancement factors (EF) are observed between 106 and 108 [104] with some reporting EFs as high as 1014, thereby indicating that single molecule detection is possible [99], [107]. The dramatic enhancement seen for SERS signals makes it useful for detecting extremely low analyte concentrations. Various groups have documented the success of SERS for detecting analytes concentrations of nano-grams per milliliter. Some have shown successful analyte detection at pico-gram per milliliter concentrations or even claim single molecule detection [96], [103], [108].

3.1 Overview of SERS toward POC monitoring

Given the potential advantages for the SERS approach listed above, several groups have published research documenting biomarker or analyte detection using SERS platforms ultimately aiming toward improving a POC technology [101]. SERS approaches have been developed for targeting tumors [109], diagnosing malaria [21], [27], identifying bacterial meningitis [110], and many other applications. This section will provide an overview of several biotechnologies that use SERS with a focus on colloidal particles. Benefits and limitations of each approach will also be presented in the interest of the future development POC technology.

3.2 Nanoparticles as SERS substrates

Starting from the 1970s, colloidal nanoparticle suspensions have served as one of the most popular SERS substrates, due to their relatively basic synthesis process requiring only a silver salt and a reducing agent to produce a metallic sol, with submicromolar detection limits. One of the first examples was demonstrated by Lee and Meisel using silver nanoparticles and carbocyanine dyes in 1982 [111]. In this work, they also observed the extra enhancements achievable when combining SERS with resonance Raman (RR) as was seen by Albrecht and Creighton using rhodamine in 1977 [112]. It was observed that by choosing analytes with molecular absorption near the excitation wavelength, as is common for many fluorophores and cell staining dyes, additional Raman signal enhancement occurs. When the resonant dye molecule is bound to a plasmonic material, both the electromagnetic and chemical enhancement mechanisms discussed previously occur and are coupled; the resulting technique is referred to as surface enhanced RR spectroscopy (SERRS). Thus, maximal signal enhancements are achieved by tailoring the resonance of the metallic substrate with that of a chromophore that is also resonant with the excitation light and capable of binding directly to the particle surface.

Not long after in 1989, Rohr, Cotton et al. revealed the first SERRS immunoassay capable of monitoring antibody/antigen interactions for the biomarker human thyroid stimulating hormone (TSH) [113]. Their one-step, no wash, sandwich-type assay used a resonant dye to indirectly monitor TSH in the physiological range, thus validating the potential of SERS to enhance and improve diagnostic assays. A decade and a half later, Nie and Emory were able to achieve SERRS enhancements of up to 1015 when probing individual rhodamine 6G molecules on single silver nanoparticles [114], thus enticing future investigators to utilize this technique for POC applications that require ultra-low LODs.

Many early approaches used planar arrays of plasmonic nano-rough structures to facilitate the translation of SERS to a widely accepted, commercially viable platform diagnostic technology. Planar SERS substrates that provide maximum enhancements by forming nanogaps between metallic particles, where plasmons can couple and generate defined “hotspots” [115]. These hot spots have a strong spatial and local field dependency that tend to occur within aggregates, at sharp edges, and areas of large curvature of metallic nanostructures (on the order of 10–100 nm) [116], [117], [118]. Due to the difficulty in reproducibly preparing nanostructures, many planar SERS substrates must be synthesized using a variety of sequential complex photolithographic techniques borrowed from the silicon wafer industry. These methods include techniques such as using electron beam deposition (EBD) to create the nanorough surface itself [90], [119], using EBD to produce “nanostamps” [120], combining nanoimprint and copolymer lithographic techniques [121], using self-assembled nanoparticles (SAMs) [122], [123], and many others using non-spherical particles such as Au- and Ag-coated nanorod arrays [124], [125]. These techniques, specifically EBD, produce high-resolution nanostructures that are reproducible within a wafer; however, there is a tradeoff with high reproducibility and low enhancement factor (102–104) [126]. Due to the difficulty of fabricating well-controlled small gaps or complex geometries, nanomaterials have also been cast into thin films [127], [128], [129] or deposited onto two-dimensional nanostructured arrays [130], [131] to create hybrid substrates and improve the efficiency and density of SERS actives sites.

However, an ideal SERS substrate should not only deliver maximal enhancement but also provide a reproducible and uniform SERS response, have a stable shelf-life, and be easily fabricated [132], [133], [134]. Although planar SERS substrates are commercially available, scanning electron microscopy/transmission electron microscopy images provided by distributers often reveal a lack of batch to batch reproducibility, making the formation of clinical calibration curves very difficult. Additionally, due to the expensive nature of using a clean room in order to produce reproducible planar SERS substrates, each individual test would cost far more than the ~$1–10 limit for POC technologies. Therefore, the focus in the following sections is on SERS substrates that utilize colloidal plasmonic nanoparticles, as they are arguably the simpler SERS substrate to produce without the need for complex equipment or extensive training.

3.2.1 Developing colloidal SERS substrates

Colloidal SERS is most commonly achieved using silver and/or gold nanoparticles between 20 and 200 nm, as they exhibit unique and tunable optical properties to facilitate the plasmonic coupling event required in SERS sensing. The nanoparticle size [135], shape [136], dispersant [137], and particularly the dielectric properties of the metal all strongly effect a colloid’s extinction spectra and SERS capabilities. To ensure a colloid’s SERS enhancement factor is uniform throughout the suspension, it is also required that the nanoparticles be somewhat monodisperse (PDI<0.300) [138]. Spheroids exhibit LSPRs that are typically within a ±120-nm window of the most commonly used laser sources: 532 nm, 633 nm, and 785 nm (Figure 1 [139]). Commonly, SERS enhancements are achieved by tuning the Raman excitation wavelength close to the intrinsic LSPR peak of the colloid or by causing the colloid’s LSPR to shift into resonance with the laser by altering the particle’s properties [140]. This has been realized using controlled salt-based aggregation, mechanical trapping or centrifugation of particles, creation of core-shell particles, or binding event that results in nanoparticle assembly formation, for example, using DNA hybridization to template core-satellite formation as described extensively by Mirkin and colleagues [141], [142], [143].

Figure 1:

Photographs (A) and extinction spectra (B) of various 60-nm metal nanoparticles (AgNPs, AuNPs, and Au/Ag nanoshells) in water. LSPR peaks are observed at 430 nm, 540 nm, and 630 nm, respectively. Reprinted with permission from [139]. Copyright 2013 Royal Society of Chemistry.

3.2.2 Label-free colloidal SERS

As Raman provides a chemical fingerprint of the probed sample, it may seem intuitive to detect disease identifying changes in an isolated biomarker’s conformational structure by looking at the Raman modes coming from the analyte itself. However, the specificity or ability to uniquely measure a desired biomarker in complex biological samples is a key factor for translating SERS technologies to the POC and could inhibit simply looking at the analyte itself. Thus, for label-free methods such as monitoring the intrinsic spontaneous Raman (SR) or stimulating the resonance Raman modes of the analyte (Figure 2), the biomarker must be a pure sample and be isolated from the complex biological sample using extraction techniques such as HPLC or an immunoassay, or the biomarker itself must exhibit vibrational modes that can be uniquely pulled out from the modes inherent in the background media [145]. Bringing these purification steps to the POC often requires immobilization of affinity ligands such as antibodies or DNA aptamers onto well plates, scintillation vials, or glass slides using basic click chemistry [146]. The downside of these ELISA or extraction kit-style designs is the complexity (i.e. not user-friendly by the WHO-defined “ASSURED” criteria) including required user intervention for washing steps to remove optically or biochemically interferent components of the sample prior to SERS analysis.

Figure 2:

(A) Schematic of a label-free assay utilizing silver colloidal nanoparticles for the detection of proteins via electrostatic aggregation of Ag NPs in the presence of the protein. (B) Example of Raman spectrum of serum albumin using label-free colloidal SERS in the detection of colorectal cancer. (C) Scatter plot of the linear discriminant scores belonging to the normal and colorectal cancer group, calculated from the datasets of albumin. Reprinted with copyright permission from [144], Copyright 2014 Journal of Photonics SPIE.

For example, our group was able to observe structure changes in the SERS spectra of the biomarker β-amyloid absorbed onto aggregated gold nanospheres [147] in a pure sample that could be suggestive of Alzheimer’s disease. A second example was conducted first by Feng et al. [148] and later Wang et al. [144] using two abundant serum proteins, albumin (Figure 2) and globulin, to detect colorectal cancer. In Wang et al.’s work, the two serum proteins had to first be purified from over 200 healthy and cancerous human serum samples. Protein samples were then added directly to hydroxylamine silver nanoparticles, and acetic acid was used to aggregate silver nanoparticles to increase the magnitude of the SERS enhancement. SERS bands were assigned to verify specific biomolecular contents of the proteins and to predict protein secondary structural changes that occur with colorectal cancer progression using the difference of the SERS spectra between healthy and cancerous samples. Principal component analysis and linear discriminant analysis were required to be used to assess the capability of this approach for identifying colorectal cancer, demonstrating a diagnostic accuracy of 100% for albumin monitoring and 99.5% for globulin. Additionally, both the albumin and globulin partial lease squares (PLS) models successfully predicted the unidentified subjects with a diagnostic accuracy of 93.5%.

These results suggest that SERS analysis of serum proteins have the potential to be a sensitive and clinically powerful means for disease detection. However, simple direct sensing efforts like these still struggle to fully translate to the POC, as they require complex sample preparation to be performed before SERS analysis. Thus, these methods are not user friendly as noted above, are time consuming, involve complex statistical analysis or peak assignments, and require too many separate pieces of laboratory equipment to be fully implemented at the patient bedside.

3.3 Molecularly mediated colloidal SERS

Indirect sensing using an assay whose SERS response is facilitated by a molecular binding event, particularly one that involves the biomarker itself, has emerged as an efficient approach to colloidal SERS. Specifically, SERS-active nanoprobes have the potential to enhance specificity [149]. In general, they can involve one or more nanoparticles conjugated to (1) a highly polarizable Raman reporter molecule, (2) an affinity ligand such as an aptamer or antibody, and (3) a steric or electrostatic capping agent for stabilization in high salt environments [150], [151].

Oligonucleotides, antibodies, protein antigens, small molecules, and dyes can all be immobilized onto metallic nanoparticles using thiol end groups, bifunctional PEG linkers, or sequential click chemistry [152], [153], [154], [155]. Extensive work has been done by groups such as Graham et al. who utilize various oligonucleotide and resonant dye-coated nanoprobes to form SERRS active nanoassembly complexes for multiplexed DNA detection [156]. In most cases, the nanoassembly detection modality involves the SERRS active particles’ LSPR shifting in or out of resonance with the excitation source. This is caused by hybridized DNA linking of the nanoparticles in close proximity in order to share conduction band electrons and red-shift their extinction spectra. They, thereby, jump in SERS intensity, without causing irreversible aggregation (Figure 3). The group has translated their “SERS-on” techniques for a variety of DNA, protein, and small molecule sensing applications, most recently by Mabbott et al. for monitoring four fungal probes in a multiplexed fashion [157], by Simpson et al. using a biomimetic glyconanoparticle assay for ultrasensitive (ng/ml) quantification of cholera toxin B-subunit [158], and by Gracie et al. for the simultaneous detection of two meningitis bacterial DNA biomarkers extracted from cerebral spinal fluid (CSF) clinical samples [110], [159].

Figure 3:

The top scheme shows the hybridization of nanoparticle probes to complementary single stranded DNA. The length of the probe sequence can be varied to increase discrimination and stability. The Raman reporter (dye) is added to the nanoparticle surface along with the DNA probe sequence to create a unique code for this particular DNA sequence. Schematics (A–C) represent the three different orientations that the probes can take when attaching to a complementary single strand of DNA due to the stereochemistry induced by the deoxyribose sugar on DNA. (A) Tail-to-tail, (B) head-to-tail, (C) head-to-head. Adapted with permission from [156]. Copyright 2015 American Chemical Society.

Another approach uses only one colloid without aggregation, relying solely on small molecule binding to either competitively displace an aptamer tagged with a Raman dye molecule in a “SERS-off” configuration (Figure 4). Chung et al. utilized this “SERS-off” molecularly mediated SERS methodology and used a partial complimentary sequence to immobilize an ssDNA aptamer onto Au/Ag core-shell nanoparticles. This method proved to be sensitive down to the 10-fm range for BPA-spiked tap water, over a total dynamic range of 10 fm–100 nm [160]. The authors of this work acknowledge that this LOD is two or three orders of magnitude lower than that reported for other BPA sensing techniques but may possibly be higher if the samples were in complex biological media. It is noteworthy that the total detection time was estimated to be about 40 min including both the reaction between aptamer and BPA (30 min) and detection (10 min), making this option good for supplementing rapid diagnostic tests (RDTs).

Figure 4:

SERS spectra for (A) cy3-labeled aptamer double strand DNA-embedded Au/Ag core–shell NPs and (B) aptamer-detached Au/Ag core-shell NPs in the presence of 100 nm BPA. Adapted with permission from [160]. Copyright 2015 Elsevier.

3.3.1 Magnetic approaches for colloidal SERS monitoring in complex samples

One obstacle preventing the translation of molecular diagnostics using SERS at the POC is the lack of simple methods that can be integrated into portable platforms, for instance, without the need for complexity such as sample washing steps. One potential technology to overcome this challenge is the use of magnetic approaches including microbeads and superparamagnetic nanoparticles (SPIONS) [161]. These are easy to manipulate with small permanent neodymium magnets held at the side of a vial, well plates, capillary tube, microfluidic channels, or even inside cells [162]. When functionalized with sensing ligands, this allows for faster, more automated washing steps while also preventing sample sedimentation often seen with repeated centrifugation [29], [163]. Magnetic nanoparticles can also provide a plasmonic response when coated in gold or silver [164], therefore improving SERS enhancement capabilities [165], [166], [167].

Many groups have facilitated this technique for improving clinical chemistry techniques recently, such as Wang et al. who used aptamers immobilized onto silver-coated magnetic nanoparticles along with a secondary SERS active gold nanoprobe coated with another aptamer to capture and quantify bacterial cells down to 10 cells/ml [168]. Ge et al. used a similar sandwich binding approach but with antibodies in place of aptamers for the detection of the ovarian cancer serum biomarker human epididymis protein 4 (HE4). They not only were able to demonstrate fg/ml LODs and a dynamic range of 1 pg/ml to 10 ng/ml but also demonstrated that the assay particles could be washed and reused at least five times in their efforts towards developing easy to use diagnostic kits [169].

Tuan Vo-Dihn’s group has also developed a sandwich-type SERS assay, relying on specific DNA hybridization to capture ultrabright SERS nanorattles onto magnetic microbeads [27]. As shown in Figure 5, nanorattles are core-shell silver particles with RR reporters loaded in the gap space between the core and the shell. The DNA probes are coated on the shell surface, thus acting as the SERS tags for signal detection. After hybridization, a magnet was applied to the bottom of the well to both remove unbound nanorattles and to concentrate the hybridization sandwiches at a localized detection area for SERS measurements. Probing for two specific DNA sequences of the malaria parasite Plasmodium falciparum, one mutated and the other wild type, it was found that SERS could detect malaria DNA down to 100 am. As the mutant sequence translates for resistance to artemisinin drugs, single nucleotide polymorphism (SNP) discrimination of wild-type malaria DNA and mutant malaria DNA was also demonstrated. Their results show the potential for molecularly mediated SERS to differentiate small mutations in infectious pathogens with far greater sensitivity than current methods, an important factor for global health applications.

Figure 5:

(A) The nanorattle-based DNA detection method using sandwich hybridization of (1) magnetic bead that are loaded with capture probes, (2) target sequence, and (3) ultrabright SERS nanorattles that are loaded with reporter probes. (B) A magnet is applied to concentrate the hybridization sandwiches at a detection spot for SERS measurements. Reprinted with permission from [27]. Copyright 2016 Elsevier.

Another relevant magnetic-based method for potential use in SERS sensing was demonstrated by He, Li, and Hu with an aptamer recognition-induced target-bridged SERS assay based on magnetic chitosan (MCS) and silver/chitosan nanoparticle (Ag@CS NPs) binding [170]. A single aptamer target nanoparticle was used for the detection of three different types of protein, benefiting from the highly specific affinity of aptamers and biocompatibility of chitosan (CS). MCS coated in various antibodies or aptamer act as capture probes in the triple sandwich assay format shown. The sandwich complexes of aptamer (antibody)/protein/aptamer were first mixed with complex biological mediate and separated from biological samples after the reaction proceeded by magnetic manipulation with a permanent magnet under a glass slide. SERS signals were collected after washing the complexes, and the protein concentrations indirectly correlated with the number of Raman report molecules left after washing. To demonstrate the translatability of this method, three different proteins – thrombin, platelet derived growth factor (PDGF), and immunoglobulin E (lgE) – were investigated. The CS shell demonstrated enhanced stability for longer shelf life and prevention of signal drift due to loss of Raman reporter.

Like many colloidal nanoparticle assays, this method avoids slow diffusion limited kinetics problems observed for solid SERS substrates. The feasibility of this method for potential use at the POC was shown with PDGF BB in clinical serum samples, with an LOD of 3.2 pg/ml. The prediction results obtained from human serum of healthy patients vs. cancer patients using the proposed SERS method correlated with traditional ELISA results while the SERS method expanded the linear range.

3.3.2 Combining SERS-based immunoassays and ELISA

SERS nanoprobes have recently been used to improve detection capabilities of immunoassays and have the potential to rival the popular enzyme-linked immunosorbent assay (ELISA) techniques. Combining SERS and ELISA, aka “SLISA”, has proven to be an effective method for improving the LODs due to the intrinsic enhancement capabilities of SERS, the ability to speed up the assay reaction times due to the 3D architecture of functionalized colloidal nanoparticles, and the ability to capitalize on the narrow spectra bands obtained with Raman for improving the multiplexing capabilities of traditional immunoassays [171]. Bhardwaj et al. directly compared the capabilities of SLISA and ELISA assays for the measuring RAD54 stress-marker proteins. They found that SLISA has similar accuracy as ELISA but improves upon the indirect enzyme-based method by being reusable, faster, more direct, and easier to use. SLISA was also five times more sensitive than ELISA while providing qualitative information on the immunosensor’s chemical characterization and antigen-antibody binding. This allows direct detection with less uncertainty, which is a stringent limitation of all label-based biosensor technologies, including ELISA [172].

One example of a biomarker candidate for SLISA is the hormone estradiol (17β-estradiol, E2), a critical serum protein in sexual development. The E2 levels are especially low (<10 pg/ml) in prepubertal girls, and current clinical detection methods are insufficient for accurate assessment of E2 at these ultralow concentrations. In a study conducted by Choo’s group, a new E2 sensor was introduced using a magnetic capture bead SERS immunoassay detection platform [87]. The work was based on their previous work that validated the technique for use with clinical samples for the early diagnosis of arthritis [173].

The system involves a competitive binding assay with reagents immobilized onto magnetic beads to assist with automated wash steps and also to enhance the SERS response through magnetic aggregation in a glass capillary tube (Figure 6). Their SERS assay was tested with 30 blood samples to assess its clinical feasibility, and their prediction results were compared to those obtained using a commercially available chemiluminescence immunoassay. The commercial immunoassay failed to quantify E2 serum levels lower than 10 pg/ml, but the LOD of E2 using the novel SERS-based assay described in this study was an order of magnitude lower at 0.65 pg/ml. This verified that SLISA-based methods have a strong potential in the early identification of biomarkers due to their exceptional analytical sensitivity.

Figure 6:

Schematic illustration of the SERS-based competitive immunoassay for quantification of E2- target where E2 and E2-conjugated SERS nanotags competitively react with anti-E2 antibody on magnetic beads. Reprinted with permission from [87]. Copyright 2016 American Chemical Society.

3.3.3 Dual modality colloidal SERS

Beyond SLISA, another emerging trend in colloidal SERS is the utilization of dual optical modality approaches. For example, many colloidal SERS assays also intrinsically exhibit a colorimetric response and dual sensing can facilitate simple yes/no readouts [174], [175]. Researchers have also combined SERS with fluorescence to provide additional visual confirmation of binding results in a multiplexed format [176]. As more methodologies emerge and combine, the benefits of SERS will only be expended even further.

4 Implementation of SERS POC technology using different fluidics platforms

The aforementioned advancement in SERS assays for the detection of biological analytes in complex media has supported its potential use in POC platforms for the detection and monitoring of different diseases. This section provides an overview of the two major platforms, microfluidics and paper-based fluidics. Both are being invested by several groups in the transformation of SERS assays towards future POC biotechnologies.

4.1 SERS combined with microfluidics

Microfluidics is the science and technology of manipulating and controlling fluids typically in the range of microliters to picoliters using microchannels [177]. The use of microfluidics for analytical biosensing has the potential to not only facilitate the assay procedure but also improve assay results [107], [177]. In particular, advances in the microfluidics technology field have contributed to the development of LOC biosensors. The use of microchannels, microvalves, micomixers, and micropumps has allowed the creation of small chips that can potentially perform all of the functions needed in an immunoassay procedure [177]. The main advantages of microfluidic based biosensors are that they can measure minimal sample volumes and potentially eliminate the need of user input in the process. Also, they usually have short times for analysis, which could be very important for POC diagnostics.

In common immunoassays procedures, the user usually has to dispose of the samples, add reagents, wash wells, mix solutions, and take measurements. The use of microfluidic based biosensors can eliminate many of these steps and thus reduce the possibility that human error can affect the measurements. This can potentially be translated into improved sensitivity, precision, ease of use, rapid results, and minimal amount of sample needed [178].

Several groups and multiple reviews have incorporated microfluidic technology with SERS-based assays to create sensitive sensors for potential POC applications [107], [178], [179], [180].

4.1.1 Mixing in the microchannel

The ability to reproducibly mix the components of an assay is essential for the appropriate functioning of a diagnostic test. The SERS-based assays that use functionalized colloidal nanoparticles typically have to be thoroughly mixed with the analyte of interest and other components to allow them react and interact to produce accurate results. Therefore, one focus in the development of a microfluidic SERS-based assay is the mixing section.

For example, Chon et al. developed a SERS-based microfluidic sensor that serially dilutes the target marker, mixes antibody-conjugated hollow gold nanospheres (HGNs) and magnetic beads, and traps the magnetic complexes with different structures. In the microchannel, a groove-shaped mixer was incorporated to improve the mixing efficiency [181]. Figure 7 shows the microfluidics design.

Figure 7:

Layout of SERS-based gradient optofluidic sensor integrated with solenoids and various regions of microchannels designed for mixing. Adapted with permission from [181]. Copyright 2010 American Chemical Society.

Wilson et al. described the development of a microchannel that uses a mixer to enhance the contact between silver colloid and an analyte. They were able to detect it with a sensitivity that was an order of magnitude greater than without using the chip [182]. In another approach, Quang et al. created a microchannel with a micropillar array to achieve efficient mixing and produce reproducible SERS detection. This microchannel allowed the detection of dipicolinin acid (DPA) and malachite green (MG) with estimated detection limits of 200 ppb and 500 ppb, respectively [183].

In another example, Geo et al. described a microfluidic based biosensor to detect the prostate-specific antigen (PSA) cancer biomarker [30]. They created a SERS-based magnetic immunoassay on a microfluidic chip, as can be observed in Figure 8. The microfluidic channel was designed to generate and mix microdroplets with the assay reagents. Inside the droplet, the antigen and the antibodies reacted to form sandwich immunocomplexes.

Figure 8:

(A) Optical image of the entire microdroplet channel filled with red ink. (B) Sequential droplet splitting mechanism for the wash-free magnetic immunoassay of PSA cancer markers: (i) mixing of reagents, (ii) formation of immunocomplexes, (iii) isolation of magnetic immunocomplexes, (iv) supernatant including unbound SERS nanotags and (v) separated magnetic immunocomplexes. Reprinted with permission from [30] Copyright 2016 American Chemical Society.

A magnetic bar embedded on the channel separated the magnetic immunocomplexes from the SERS nanotags unbound to the magnetic beads. The droplet was split into two parts with Y-shaped channel bifurcation. The fission created two droplets, one with the magnetic immunocomplexes and the second one with the unbound SERS nanotags. The SERS signal of the droplet containing the SERS nanotags was measured and analyzed. The LOD of the SERS-based microdroplet sensor was estimated to be below 1 ng/ml, which is lower than the value used in common diagnostics of PSA [30].

4.1.2 Nanoparticle aggregation and SERS substrates in microchannels

SERS can be used to develop specific and sensitive biosensors. However, a main challenge of this modality is to obtain reproducible results from measurements. The reproducibility of SERS measurements is affected by different factors such as the type of substrate used, the aggregation method, and the inhomogeneous distribution of molecules on the metallic substrate [107]. Thus, controlling the aggregation of colloid used in SERS sensors has been a main focus, as the development of this technology. A reproducible aggregation of nanoparticles or controlled deposition of enhancement structures on a surface can be translated into a consistent SERS enhancement. As a result, several groups have tried different approaches to control the aggregation of nanoparticles or to produce reproducible SERS substrates on surfaces with defined enhancement spots [107].

Wang et al. developed an optofluidic device with a microchannel-nanochannel junction to trap and assemble nanoparticles into SERS active clusters by using capillary force (Figure 9). This cluster provided an electromagnetic enhancement factor of about 108. However, they reported a SERS enhancement reproducibility of ±10% (device to device) when 83 nm of adenine was used [184].

Figure 9:

Schematic diagram of an optofluidic device: (A) side view, (B) top view, (C) side view of an optofluidic device with aggregated nanoparticle-SERS active clusters at the step structure. The depth of nanochannel is smaller than that of nanoparticles. Thus, nanoparticles are trapped and aggregated. (D) Fluorescent image of polystyrene nanoparticles trapped at the step boundary of the optofluidic device. Reprinted with permission from [184]. Copyright 2007 American Chemical Society.

Yazdi and White described another aggregation method by forming a 3D nanofluidic network with packed nanoporous silica microspheres in a microfluidic channel [185]. This matrix trapped silver nanoclusters and adsorbed analytes into the SERS detection area. With this approach, a concentration of R6G of 400 am was detected. A multimode fiber optic was also integrated in the channel, which eliminated the need for optical alignment.

Magnetic sections have also been used to aggregate nanoparticles for SERS measurement. Gao et al. demonstrated an assay to detect the anthrax biomarker poly-y-D-glutamic acid (PGA) on a microfluidic chip (Figure 10). In this assay, PGA and PGA-conjugated gold nanoparticles competed for binding sites on anti-PGA-immobilized magnetic beads. The magnetic immunocomplexes were trapped by yoke-type-solenoids embedded on the microchannel where the SERS signals were measured. The assay estimated LOD was 100 pg/ml [186].

Figure 10:

(A) Schematic illustration of the solenoid-embedded dual channel microfluidic sensor. The sensor is composed of two parallel channels: one for PGA sensing (light gray) and the other for control (dark gray). (B) Optical images of the solenoid chip filled with four different colors of inks. (C) Photograph of the capture area for magnetic immunocomplexes. Reprinted with permission from [186]. Copyright 2015 American Chemical Society.

Instead of aggregating nanoparticles to create the SERS enhancement, other groups have developed microfluidic devices with nano-rough substrates already incorporated. For example, Liu et al. described the creation of nanowell structures on PDMS. A thin Ag film was deposited on the nanowells to create SERS active sites. The SERS spectra of Rhodamine 6G and adenosine were measured with the microchannel. The SERS enhancement on the nanowell-based Ag SERS substrate was more than 107 times higher than on a smooth Ag layer on PDMS [187]. A similar microchannel was reported by Oh et al. where plasmonic nanoprobes with hotspots were selectively patterned on PDMS microchannels. This microchannel enabled solution-phase SERS detection of small molecules [188].

Most microfluidic devices are made from polydimethylsiloxane (PDMS) because the fabrication of channel systems is straightforward and the channels are flexible. PDMS channels can also be combined with SERS substrates prepared on a surface, such as glass [187].

As mentioned, the advantages of using microfluidics for potential POC diagnosis include the use of minimal volumes; the ability to control the sample and perform steps such as mixing and washing automatically; the ability to increase the SERS substrate reproducibility, which improves precision; and the ability to rapidly detect biomarkers with high sensitivity. However, some of the disadvantages of microfluidic based sensors can be the cost to manufacture the chips and, in some cases, the need for external devices to control fluid flow. Thus, new technologies such as paper-based SERS biosensors are being developed to address these issues.

4.2 Paper-based SERS platforms

There are several different fabrication techniques for paper-based sensing microfluidics reported in the literature [189], [190], [191], [192]. However, for this review, only techniques utilized in combination with SERS-based detection towards POC will be discussed.

Paper-based SERS is attractive due to its low cost, simplicity, ability for multiplexing, and reduction in analysis time. Typically, these “paper fluidics” are cellulose based, allowing for the flow or imprinting of nanoparticles embedded within its matrix. This technique has been adapted from chromatography where samples can be separated and analyzed based on size, shape, or surface charge, which allows for separation, detection, and analysis, all on a single platform. This section focuses on different fabrication techniques and monitoring approaches for paper-based SERS detection that have the potential to be adopted for POC applications.

4.2.1 Filter membrane-based SERS with a syringe

Filter membrane-based SERS detection utilizes inexpensive tools such as a disposable syringe, filter holder, and filter membrane. The basic concept involves pre-wetting the filter membrane with an organic solvent such as ethanol, passing colloidal (Au or Ag)-coated nanoparticles through the filter membrane via the syringe and then passing the sample through the filter (Figure 11) [193]. The colloidal particles are coated with the desired target probe that interacts with the sample. The filter membrane allows for the aggregation of these particles for SERS detection via portable spectrometer after removal and drying of the membrane. Research groups have utilized this quick and easy method for detection of chemical and biological entities such as melamine and malathion [193], Escherichia coli [194], and other toxins or pathogens in food [195]. Due to the simplicity of this approach, the technique can be utilized in on-site diagnosis. However, White’s group reported that this technique was shown to be two to three orders of magnitude less sensitive than other paper-based SERS methods [193].

Figure 11:

SERS-active substrates are created simply by passing a silver colloid solution through a filter membrane using a syringe. Analyte molecules are concentrated into the substrate from a large sample volume. The SERS signal is detected using a small and portable photonic setup. Reprinted with permission from [193]. Copyright 2012 American Chemical Society.

4.2.2 Dip coating paper-based SERS

Filter paper can also be used to absorb nanoparticles via immersion of the filter paper into solutions of colloidal nanoparticles. Typically, filter paper such as Whatman® is submerged for a period of time in solution of colloidal nanoparticles. The paper is then removed and allowed to air dry. The particles are retained within the filter’s matrix via adsorption due to van der Waals forces and hydrophobic interactions between the nanoparticles and fibers [196]. Groups such as Cheng et al., Liu et al., and Ngo et al. have utilized this technique to detect tyrosine [197], oral cancer cells [198], and antigen [199], respectively. Cheng et al. reported that the use of filter paper to detect tyrosine provided 50 times more SERS signal and a detection limit of 625 nm [197]. This approach provides an easy fabrication method of the SERS substrate and can be widely used as a swab or dipstick to collect samples.

4.2.3 Printed paper-based SERS substrates

Commercially available inkjet printers can be reengineered for use in printing highly concentration colloidal nanoparticles on specific regions of the paper or for patterning hydrophobic barriers to direct the flow of the fluid. Typically, either a thermal or a piezoelectric printer is used depending on the thermal sensitivity of the SERS substrate and/or the solvent being used.

In terms of nanoparticle printing, Yu and White demonstrated that colloidal silver nanoparticle arrays can be printed on cellulose based paper to form aggregates within its matrix using an EPSON Workforce 30 inkjet printer [200]. The printed paper was then cut into a specific shape and dipped into a solution containing the analyte of interest. After time was given for the liquid to be wicked and allowed to travel to the assay region, a fiber optic, portable spectrometer (excitation wavelength of 785 nm and laser power of 17 mW) was used to detect varying concentrations of drugs and pesticides on the nanogram scale [200], [201]. This technique provided high density aggregation of the colloidal nanoparticles to enhance the SERS signal. However, this printing approach can be difficult to produce repeatedly. Other limitations also include complications in modification of the printer and problems with nozzle clogging.

Besides using the printer to deposit nanoparticles on the surface of paper, the high resolution of inkjet printing can also be used to pattern hydrophobic borders on hydrophilic paper to create microchannels. These hydrophilic channels created within the paper are used to direct the flow of fluids through the membrane to regions of assay interaction and sensing. This is typically accomplished using a patterning agent such as wax and selectively creating hydrophobic regions. Torul et al. demonstrated the detection of glucose from whole blood by placing a droplet of blood on a gold wax printed paper containing gold nanorods [202].

SERS substrates can be screen printed on filter paper at an even lower cost than ink-jet printing. This process is carried out by forcing a high concentration of colloidal nanoparticles through a meshed screen printing plate designed with a desired patterned aperture [203]. This creates multiple areas of SERS reactive regions on the paper. After drying, the paper-based SERS substrate can then be exposed to various samples and probed with a portable Raman spectrometer for analyte detection. Qu et al. used this technique to detect multiple biological analyte on a single paper with LODs ranging from 10−7 to 10−10 [203]. However, one major disadvantage to using this approach is controlling the viscosity of the colloidal nanoparticles without significantly hindering the SERS signal.

4.2.4 Lateral flow paper-based SERS

One of the earliest and most widespread POC devices is based on the concept of lateral flow and used to create off the shelf testing platforms, such as the at-home pregnancy test and glucose self-testing strips [204], [205]. In recent years, this same approach has become attractive once again to create rapid detection kits for more complex biological analytes in the early stages of disease (Figure 12) [206]. Lateral flow relies on capillary forces to move small molecules along a transport medium such as cellulose paper. By combining this technique with chromatography, more complex media can be collected, separated, and analyzed all on one low-cost, easily fabricated testing platform. Lateral flow paper-based microfluidics eliminates the need for precise patterning or printing and reduces the cost of equipment because the flow and separation are dependent on the shape, size, and sharpness of the corners of the paper. Fu and Choo incorporated this technique to design a POC technology capable of detecting the human immunodeficiency virus type 1 (HIV-1) DNA [207]. Choi and Choo also demonstrated another clinical application of this approach for the detection of the thyroid-stimulating hormone in biological fluids in diagnosing hyper/hypo-thyroidism [208]. They reported that this approach was two orders of magnitude more sensitive than conventional colorimetric approaches. Choo’s group has also applied this technology to design a SERS-based immunoassay for staphylococcal enterotoxin B [206].

Figure 12:

Schematic of a lateral flow based platform for the detection of analytes from whole blood using paper chromatography. (a) Qualitative analysis; (b) Quantitative analysis. Reprinted with permission from [206]. Copyright 2016 American Chemical Society.

4.2.5 Current limitations of paper-based SERS microfluidics

Overall, printed paper microfluidics for SERS detection of chemical and biological analytes is an attractive technology because it offers an affordable and easy method to mass produce devices that can be used with a portable Raman spectrometer for potential on-site diagnosis. However, many of the paper-based approaches described are reasonable for detection but not yet adequate for quantification. In other words, they have been used primarily to determine whether or not the biomarker is present but rarely used to determine the repeatable concentration of the biomarker across a dynamic range. Furthermore, although the Raman spectrometers have been reduced in both size and price over the past two decades, the hand-held devices are still too expensive for delivering POC devices. If portable Raman spectrometers could be mass produced for high volume POC applications, rather than primarily used as research tools as they are now, it is reasonable to assume the cost and size will continue to be reduced.

5 Conclusions

POC technologies that follow the WHO’s “ASSURED” guideline can offer many advantages over current bench-top lab-based measurements and provide a means for RDT. In this review, SERS has been explored as a dynamic technique for POC monitoring because of its high sensitivity (up to fm detection limits) and multiplexing capabilities. Many SERS-based assays and platforms are currently under development. This review focused on comparing these systems in terms of the synthesis, functionalization, and utilization of plasmonic nanoparticles as the SERS substrates within different environments including microwells, microfluidics, and paper-based platforms. In particular, device complexity can be reduced by coupling the SERS-based substrates with low cost, easy to fabricate, paper fluidics.

While the high sensitivity and multiplexing ability of SERS hold promise for its use in POC diagnostics, it still fails to produce signals as robust and repeatable as current gold standard assays. Additionally, researchers rarely present methodologies that can be translated across multiple biomarker types or for analytical ranges spanning over several orders of magnitude. However, advances in more specific and robust capture ligands such as DNA aptamer-coated nanomaterials offer a promising emergent solution. Furthermore, SERS is also limited by the high cost of portable Raman readers although hand-held systems have been significantly reduced in size and cost over the past decade. Overall, the promise for SERS to be used for POC monitoring will rely on overcoming the barriers through further advancements in the assays, platforms, and more dedicated, cost-effective, Raman readers.


The authors wish to acknowledge the financial support of the National Institutes of Health (2R44ES022303). Furthermore, we would like to thank Samuel Mabbott for his helpful discussions and review of this manuscript.


  • [1]

    Gubala V, Harris LF, Ricco AJ, Tan MX, Williams DE. Point of care diagnostics: status and future. Anal Chem 2012;84:487–515.Google Scholar

  • [2]

    Plebani M. Harmonization in laboratory medicine: the complete picture. Clin Chem Lab Med 2013;51:741–51.Google Scholar

  • [3]

    Nichols J, Ehrmeyer S-R, Greenberg N, Mett-Stabler CAH, Master DS, Valdes R. Laboratory medicine: advancing quality in patient care. Am Assoc Clin Chem 2015. Available at: https://www.aacc.org/health-and-science-policy/aacc-policy-reports/2015/laboratory-medicine-advancing-quality-in-patient-care.Google Scholar

  • [4]

    Chan CP, Mak WC, Cheung KY, et al. Evidence-based point-of-care diagnostics: current status and emerging technologies. Annu Rev Anal Chem (Palo Alto Calif) 2013;6:191–211.Google Scholar

  • [5]

    Howick J, Cals JW, Jones C, et al. Current and future use of point-of-care tests in primary care: an international survey in Australia, Belgium, The Netherlands, the UK and the USA. Br Med J Open 2014;4:e005611.Google Scholar

  • [6]

    Kessler R, Glasgow RE. A proposal to speed translation of healthcare research into practice: dramatic change is needed. Am J Prev Med 2011;40:637–44.Google Scholar

  • [7]

    Yager P, Domingo GJ, Gerdes J. Point-of-care diagnostics for global health. Annu Rev Biomed Eng 2008;10:107–44.Google Scholar

  • [8]

    Woolley CF, Hayes MA. Emerging technologies for biomedical analysis. Analyst 2014;139:2277–88.Google Scholar

  • [9]

    Horvath AR, Lord SJ, St John A, et al. From biomarkers to medical tests: the changing landscape of test evaluation. Clin Chim Acta 2014;427:49–57.Google Scholar

  • [10]

    Sohn AJ, Hickner JM, Alem F. Use of point-of-care tests (POCTs) by US primary care physicians. J Am Board Fam Med 2016;29:371–6.Google Scholar

  • [11]

    Poste G. Bring on the biomarkers. Nature 2011;469:156–7.Google Scholar

  • [12]

    John AS, Price CP. Existing and emerging technologies for point-of-care testing. Clin Biochem Rev 2014;35:155–67.Google Scholar

  • [13]

    Granger JH, Schlotter NE, Crawford AC, Porter MD. Prospects for point-of-care pathogen diagnostics using surface-enhanced Raman scattering (SERS). Chem Soc Rev 2016;45:3865–82.Google Scholar

  • [14]

    Lopez-Barbosa N, Gamarra JD, Osma JF. The future point-of-care detection of disease and its dat a capture and handling. Anal Bioanal Chem 2016;408:2827–37.Google Scholar

  • [15]

    Garg SK, Hirsch IB. Self-monitoring of blood glucose. Diabetes Technol Ther 2016;18:S3–9.Google Scholar

  • [16]

    Tokel O, Inci F, Demirci U. Advances in plasmonic technologies for point of care applications. Chem Rev 2014;114:5728–52.Google Scholar

  • [17]

    Pohanka M, Skládal P. Electrochemical biosensors – principles and applications. J Appl Biomed 2008;6:57–64.Google Scholar

  • [18]

    Thévenot DR, Toth K, Durst RA, Wilson GS. Electrochemical biosensors: recommended definitions and classification. Anal Lett 2001;34:635–59.Google Scholar

  • [19]

    Wang J. Electrochemical biosensors: towards point-of-care cancer diagnostics. Biosens Bioelectron 2006;21:1887–92.Google Scholar

  • [20]

    Mehrvar M, Abdi M. Recent developments, characteristics, and potential applications of electrochemical biosensors. Anal Sci 2004;20:1113–26.Google Scholar

  • [21]

    Liu Q, Yuen C. Effect of magnetic field in malaria diagnosis using magnetic nanoparticles. In European Conference on Biomedical Optics 2011;8087:80870E 1–4.Google Scholar

  • [22]

    Park HY, Schadt MJ, Wang L, et al. Fabrication of magnetic core@Shell Fe oxide@Au nanoparticles for interfacial bioactivity and bio-separation. Langmuir 2007;23:9050–6.Google Scholar

  • [23]

    Gijs MAM. Magnetic bead handling on-chip: new opportunities for analytical applications. Microfluid Nanofluidics 2004;1:22–40.Google Scholar

  • [24]

    Wang Z, Bai Y, Wei W, Xia N, Du Y. Magnetic Fe3O4-based sandwich-type biosensor using modified gold nanoparticles as colorimetric probes for the detection of dopamine. Materials 2013;6:5690–9.Google Scholar

  • [25]

    Ajroudi L, Mliki N, Bessais L, Madigou V, Villain S, Leroux C. Magnetic, electric and thermal properties of cobalt ferrite nanoparticles. Mater Res Bull 2014;59:49–58.Google Scholar

  • [26]

    Zhang J, Joshi P, Zhou Y, Ding R, Zhang P. Quantitative SERS-based DNA detection assisted by magnetic microspheres. Chem Commun 2015;51:15284–6.Google Scholar

  • [27]

    Ngo HT, Gandra N, Fales AM, Taylor SM, Vo-Dinh T. Sensitive DNA detection and SNP discrimination using ultrabright SERS nanorattles and magnetic beads for malaria diagnostics. Biosens Bioelectron 2016;81:8–14.Google Scholar

  • [28]

    Donnelly T, Smith WE, Faulds K, Graham D. Silver and magnetic nanoparticles for sensitive DNA detection by SERS. Chem Commun 2014;50:12907–10.Google Scholar

  • [29]

    Tekin HC, Gijs MA. Ultrasensitive protein detection: a case for microfluidic magnetic bead-based assays. Lab Chip 2013;13:4711–39.Google Scholar

  • [30]

    Gao R, Cheng Z, deMello AJ, Choo J. Wash-free magnetic immunoassay of the PSA cancer marker using SERS and droplet microfluidics. Lab Chip 2016;16:1022–9.Google Scholar

  • [31]

    Johnson BN, Mutharasan R. Biosensor-based microRNA detection: techniques, design, performance, and challenges. Analyst 2014;139:1576–88.Google Scholar

  • [32]

    Myers FB, Lee LP. Innovations in optical microfluidic technologies for point-of-care diagnostics. Lab Chip 2008;8:2015–31.Google Scholar

  • [33]

    Borisov SM, Wolfbeis OS. Optical biosensors. Chem Rev 2008;108:423–61.Google Scholar

  • [34]

    Ligler FS. Perspective on optical biosensors and integrated sensor systems. Anal Chem 2009;81:519–26.Google Scholar

  • [35]

    Xu M, Luo X, Davis JJ. The label free picomolar detection of insulin in blood serum. Biosens Bioelectron 2013;39:21–5.Google Scholar

  • [36]

    Gallegos D, Long KD, Yu H, et al. Label-free biodetection using a smartphone. Lab Chip 2013;13:2124–32.Google Scholar

  • [37]

    Chua JH, Chee R-E, Agarwal A, Wong SM, Zhang G-J. Label-free electrical detection of cardiac biomarker with complementary metal-oxide semiconductor-compatible silicon nanowire sensor arrays. Anal Chem 2009;81:6266–71.Google Scholar

  • [38]

    Fang X, Tan OK, Tse MS, Ooi EE. A label-free immunosensor for diagnosis of Dengue infection with simple electrical measurements. Biosens Bioelectron 2010;25:1137–42.Google Scholar

  • [39]

    Daniels JS, Pourmand N. Label-free impedance biosensors: opportunities and challenges. Electroanalysis 2007;19: 1239–57.Google Scholar

  • [40]

    Li M, Zhao F, Zeng J, Qi J, Lu J, Shih W-C. Microfluidic surface-enhanced Raman scattering sensor with monolithically integrated nanoporous gold disk arrays for rapid and label-free biomolecular detection. J Biomed Optics 2014;19:1116111.Google Scholar

  • [41]

    Bryan T, Luo X, Bueno PR, Davis JJ. An optimised electrochemical biosensor for the label-free detection of C-reactive protein in blood. Biosens Bioelectron 2013;39:94–8.Google Scholar

  • [42]

    Vestergaard MD, Kerman K, Tamiya E. An overview of label-free electrochemical protein sensors. Sensors 2007;7:3442–58.Google Scholar

  • [43]

    Gao Z, Agarwal A, Trigg AD, et al. Silicon nanowire arrays for label-free detection of DNA. Anal Chem 2007;79:3291–7.Google Scholar

  • [44]

    Ge S, Liu F, Liu W, Yan M, Song X, Yu J. Colorimetric assay of K-562 cells based on folic acid-conjugated porous bimetallic Pd@Au nanoparticles for point-of-care testing. Chem Commun 2014;50:475–7.Google Scholar

  • [45]

    Sato K, Hosokawa K, Maeda M. Colorimetric biosensors based on DNA-nanoparticle conjugates. Anal Sci 2007;23:17–20.Google Scholar

  • [46]

    Mazzone PJ, Hammel J, Dweik R, et al. Diagnosis of lung cancer by the analysis of exhaled breath with a colorimetric sensor array. Thorax 2007;62:565–8.Google Scholar

  • [47]

    Miao P, Liu T, Li X, Ning L, Yin J, Han K. Highly sensitive, label-free colorimetric assay of trypsin using silver nanoparticles. Biosens Bioelectron 2013;49:20–4.Google Scholar

  • [48]

    Shen L, Hagen JA, Papautsky I. Point-of-care colorimetric detection with a smartphone. Lab Chip 2012;12:4240–3.Google Scholar

  • [49]

    Safavieh M, Ahmed MU, Sokullu E, Ng A, Braescu L, Zourob M. A simple cassette as point-of-care diagnostic device for naked-eye colorimetric bacteria detection. Analyst 2014; 139:482–7.Google Scholar

  • [50]

    Locke AK, Norwood N, Marks HL, et al. Aptamer conjugated silver nanoparticles for the detection of interleukin 6, in Plasmonics in Biology and Medicine XIII, San Francisco, California, 2016, p. 972412.Google Scholar

  • [51]

    Unser S, Bruzas I, He J, Sagle L. Localized surface plasmon resonance biosensing: current challenges and approaches. Sensors 2015;15:15684–716.Google Scholar

  • [52]

    Li M, Cushing SK, Wu N. Plasmon-enhanced optical sensors: a review. Analyst 2015;140:386–406.Google Scholar

  • [53]

    Heller A, Feldman B. Electrochemical glucose sensors and their applications in diabetes management. Chem Rev 2008;108:2482–505.Google Scholar

  • [54]

    Amy Tenderich M. Use of blood glucose meters among people with Type 2 diabetes- patient perspectives. Diabetes Spectrum 2013;26:67–70.Google Scholar

  • [55]

    Adams DA, Buus-Frank M. Point-of-care technology – the i-STAT system for bedside blood analysis. J Pediatric Nursing 1995;10:194–8.Google Scholar

  • [56]

    Philips. Minicare I-20 Enabling near patient blood testing in the acute care setting, 2016. Available: http://www.philips.co.uk/healthcare/product/HCNOCTN496/minicare-i20-enabling-near-patient-blood-testing-in-the-acute-care-setting.Google Scholar

  • [57]

    On-the-spot information for when you’re on the spot, in Factsheet Inside innovation Minicare Acute Care, Philips, Ed., ed: ININ0NN, 2014.Google Scholar

  • [58]

    Vashist SK, Luppa PB, Yeo LY, Ozcan A, Luong JH. Emerging technologies for next-generation point-of-care testing. Trends Biotechnol 2015;33:692–705.Google Scholar

  • [59]

    Drain PK, Hyle EP, Noubary F, et al. Diagnostic point-of-care tests in resource-limited settings. Lancet Infect Dis 2014;14:239–49.Google Scholar

  • [60]

    Chin CD, Chin SY, Laksanasopin T, Sia SK. Low-cost microdevices for point-of-care testing. In: Issadore D, Westervelt RM, eds. Point-of-care diagnostics on a chip. New York, Springer, 2013, 3–21.Google Scholar

  • [61]

    Tomazelli CWK, Cheng CM, Carrilho E, de Jesus DP. Recent advances in low-cost microfluidic platforms for diagnostic applications. Electrophoresis 2014;35:2309–24.Google Scholar

  • [62]

    Sharma S, Zapatero-Rodriguez J, Estrela P, O’Kennedy R. Point-of-care diagnostics in low resource settings: present status and future role of microfluidics. Biosensors 2015;5:577–601.Google Scholar

  • [63]

    InnovaBiosciences. Lateral flow immunoassays, 2016. Available: https://www.innovabiosciences.com/applications/lateral-flow-immunoassays.html.Google Scholar

  • [64]

    Kasera S, Herrmann LO, del Barrio J, Baumberg JJ, Scherman OA. Quantitative multiplexing with nano-self-assemblies in SERS. Sci Rep 2014;4:6785.Google Scholar

  • [65]

    Teh YJ, Bahari Jambek A, Hashim U. A study of nano-biosensors and their output amplitude analysis algorithms. J Med Eng Technol 2016;41:72–80.Google Scholar

  • [66]

    Lazcka O, Del Campo FJ, Munoz FX. Pathogen detection: a perspective of traditional methods and biosensors. Biosens Bioelectron 2007;22:1205–17.Google Scholar

  • [67]

    Yang D, Singh A, Wu H, Kroe-Barrett R. Comparison of biosensor platforms in the evaluation of high affinity antibody-antigen binding kinetics. Anal Biochem 2016;508:78–96.Google Scholar

  • [68]

    Caliendo AM, Gilbert DN, Ginocchio CC, et al. Better tests, better care: improved diagnostics for infectious diseases. Clin Infect Dis 2013;57 Suppl 3:S139–70.Google Scholar

  • [69]

    Oh SW, Moon JD, Park SY, et al. Evaluation of fluorescence hs-CRP immunoassay for point-of-care testing. Clin Chim Acta 2005;356:172–7.Google Scholar

  • [70]

    Zhao W, Zhang WP, Zhang ZL, et al. Robust and highly sensitive fluorescence approach for point-of-care virus detection based on immunomagnetic separation. Anal Chem 2012;84:2358–65.Google Scholar

  • [71]

    Zhang RQ, Liu SL, Zhao W, et al. A simple point-of-care microfluidic immunomagnetic fluorescence assay for pathogens. Anal Chem 2013;85:2645–51.Google Scholar

  • [72]

    Qin Q-P, Peltola O, Pettersson K. Time-resolved fluorescence resonance energy transfer assay for point-of-care testing of urinary albumin. Clin Chem 2003;49:1105–13.Google Scholar

  • [73]

    Bertolin G, Sizaire F, Herbomel G, Reboutier D, Prigent C, Tramier M. A FRET biosensor reveals spatiotemporal activation and functions of aurora kinase A in living cells. Nat Commun 2016;7:12674.Google Scholar

  • [74]

    Roda A, Guardigli M, Michelini E, Mirasoli M. Bioluminescence in analytical chemistry and in vivo imaging. TrAC Trends Anal Chem 2009;28:307–22.Google Scholar

  • [75]

    Bartholomeusz DA, Andrade JD. Bioluminescent based Chemchip for point-of-care diagnostics. In: IEEES-EMBS, Lyon, France, 2000.Google Scholar

  • [76]

    Griss R, Schena A, Reymond L, et al. Bioluminescent sensor proteins for point-of-care therapeutic drug monitoring. Nat Chem Biol 2014;10:598–603.Google Scholar

  • [77]

    Hu D, Fry SR, Huang JX, et al. Comparison of surface plasmon resonance, resonant waveguide grating biosensing and enzyme linked immunosorbent assay (ELISA) in the evaluation of a dengue virus immunoassay. Biosensors 2013;3:297–311.Google Scholar

  • [78]

    Chien FC, Chen SJ. A sensitivity comparison of optical biosensors based on four different surface plasmon resonance modes. Biosens Bioelectron 2004;20:633–42.Google Scholar

  • [79]

    Cai H-H, Yang P-H, Feng J, Cai J. Immunoassay detection using functionalized gold nanoparticle probes coupled with resonance Rayleigh scattering. Sens Actuators B Chem 2009;135:603–9.Google Scholar

  • [80]

    Cao C, Sim SJ. Resonant Rayleigh light scattering response of individual Au nanoparticles to antigen-antibody interaction. Lab Chip 2009;9:1836–9.Google Scholar

  • [81]

    Neely A, Perry C, Varisli B, et al. Ultrasensitive and highly selective detection of Alzheimer’s disease biomarker using two-photon Rayleigh scattering properties of gold nanoparticle. ACSNANO 2009;3:2834–40.Google Scholar

  • [82]

    Lucas LJ, Han JH, Chesler J, Yoon JY. Latex immunoagglutination assay for a vasculitis marker in a microfluidic device using static light scattering detection. Biosens Bioelectron 2007;22:2216–22.Google Scholar

  • [83]

    Garza JT, Cote GL. Design of Raman active nanoparticles for SERS-based detection. In: Colloidal Nanoparticles for Biomedical Applications XI, San Francisco, California, 2016, 97221B.Google Scholar

  • [84]

    Benford ME, Lakowicz JR, Wang M, Kameoka J, Coté GL. Detection of cardiac biomarkers exploiting surface enhanced Raman scattering (SERS) using a nanofluidic channel based biosensor towards coronary point-of-care diagnostics. In: Plasmonics in Biology and Medicine VI, San Francisco, California, 2009, 719203.Google Scholar

  • [85]

    Hoppmann EP, Yu WW, White IM. Detection of deoxyribonucleic acid (DNA) targets using polymerase chain reaction (PCR) and paper surface-enhanced Raman spectroscopy (SERS) chromatography. Appl Spectrosc 2014;68:909–15.Google Scholar

  • [86]

    Walton B, Huang P-J, Kameoka J, Deutz N, Coté GL. Development of an optofluidic SERS-based biomedical sensor. In: Optical Diagnostics and Sensing XVI: Toward Point-of-Care Diagnostics, San Francisco, California, 2016.Google Scholar

  • [87]

    Wang R, Chon H, Lee S, et al. Highly sensitive detection of hormone estradiol E2 using surface-enhanced Raman scattering based immunoassays for the clinical diagnosis of precocious puberty. ACS Appl Mater Interf 2016;8:10665–72.Google Scholar

  • [88]

    Wu HY, Cunningham BT. Point-of-care detection and real-time monitoring of intravenously delivered drugs via tubing with an integrated SERS sensor. Nanoscale 2014;6:5162–71.Google Scholar

  • [89]

    Granger JH, Schlotter NE, Crawford AC, Porter MD. Prospects for point-of-care pathogen diagnostics using surface-enhanced Raman scattering (SERS). Chem Soc Rev 2016;45:3865–82.Google Scholar

  • [90]

    Tokel O, Inci F, Demirci U. Advances in plasmonic technologies for point of care applications. Chem Rev 2014;114:5728–52.Google Scholar

  • [91]

    Foudeh AM, Didar TF, Veres T, Tabrizian M. Microfluidic designs and techniques using lab-on-a-chip devices for pathogen detection for point-of-care diagnostics. Lab Chip 2012;12:3249–66.Google Scholar

  • [92]

    Rusling JF, Kumar CV, Gutkind JS, Patel V. Measurement of biomarker proteins for point-of-care early detection and monitoring of cancer. Analyst 2010;135:2496–511.Google Scholar

  • [93]

    Berg B, Cortazar B, Tseng D, et al. Cellphone-based hand-held microplate reader for point-of-care testing of enzyme-linked immunosorbent assays. ACS Nano 2015;9:7857–66.Google Scholar

  • [94]

    Newman J, Chen K, Leona M, Vo-Dinh T. Surface-enhanced Raman scattering for identification of organic pigments and dyes in works of art and cultural heritage material. Sensor Rev 2007;27:109–20.Google Scholar

  • [95]

    Zhang JY, Do J, Premasiri WR, Ziegler LD, Klapperich CM. Rapid point-of-care concentration of bacteria in a disposable microfluidic device using meniscus dragging effect. Lab Chip 2010;10:3265–70.Google Scholar

  • [96]

    Yazdi SH, Giles KL, White IM. Multiplexed detection of DNA sequences using a competitive displacement assay in a microfluidic SERRS-based device. Anal Chem 2013;85:10605–11.Google Scholar

  • [97]

    Varsányi G, Láng L, Kovner MA. Assignments for vibrational spectra of seven hundred benzene derivatives. In: Lang L, ed. Institute of Physics Publishing. New York, Wiley, 1974, 668.Google Scholar

  • [98]

    Butler HJ, Ashton L, Bird B, et al. Using Raman spectroscopy to characterize biological materials. Nat Protocols 2016;11:664–87.Google Scholar

  • [99]

    Bantz KC, Meyer AF, Wittenberg NJ, et al. Recent progress in SERS biosensing. Phys Chem Chem Phys 2011;13:11551–67.Google Scholar

  • [100]

    Betz JF, Yu WW, Cheng Y, White IM, Rubloff GW. Simple SERS substrates: powerful, portable, and full of potential. Phys Chem Chem Phys 2014;16:2224–39.Google Scholar

  • [101]

    Cialla D, Marz A, Bohme R, et al. Surface-enhanced Raman spectroscopy (SERS): progress and trends. Anal Bioanal Chem 2012;403:27–54.Google Scholar

  • [102]

    Yang H, Deng M, Ga S, et al. Capillary-driven surface-enhanced Raman scattering (SERS)-based microfluidic chip for abrin detection. Nanoscale Res Lett 2014;9:138.Google Scholar

  • [103]

    McNay G, Eustace D, Smith WE, Faulds K, Graham D. Surface-enhanced Raman scattering (SERS) and surface-enhanced resonance Raman scattering (SERRS): a review of applications. Appl Spectrosc 2011;65:825–37.Google Scholar

  • [104]

    Vo-Dinh T, Liu Y, Fales AM, et al. SERS nanosensors and nanoreporters: golden opportunities in biomedical applications. Wiley Interdiscip Rev Nanomed Nanobiotechnol 2015;7:17–33.Google Scholar

  • [105]

    McMahon JM, Henry A-I, Wustholz KL, et al. Gold nanoparticle dimer plasmonics: finite element method calculations of the electromagnetic enhancement to surface-enhanced Raman spectroscopy. Anal Bioanal Chem 2009;394:1819–25.Google Scholar

  • [106]

    Ross MB, Ashley MJ, Schmucker AL, et al. Structure–function relationships for surface-enhanced Raman spectroscopy-active plasmonic paper. J Phys Chem C 2016;120:20789–97.Google Scholar

  • [107]

    Chao W, Chenxu Y. Analytical characterization using surface-enhanced Raman scattering (SERS) and microfluidic sampling. Nanotechnology 2015;26:092001.Google Scholar

  • [108]

    Bell SE, Sirimuthu NM. Quantitative surface-enhanced Raman spectroscopy. Chem Soc Rev 2008;37:1012–24.Google Scholar

  • [109]

    Qian XM, Peng XH, Ansari DO, et al. In vivo tumor targeting and spectroscopic detection with surface-enhanced Raman nanoparticle tags. Nat Biotechnol 2008;26:83–90.Google Scholar

  • [110]

    Gracie K, Correa E, Mabbott S, et al. Simultaneous detection and quantification of three bacterial meningitis pathogens by SERS. Chem Sci 2014;5:1030–40.Google Scholar

  • [111]

    Lee PC, Meisel D. Adsorption and surface-enhanced Raman of dyes on silver and gold sols. J Phys Chem 1982;86:3391–5.Google Scholar

  • [112]

    Albrecht MG, Creighton JA. Anomalously intense Raman spectra of pyridine at a silver electrode. J Am Chem Soc 1977;99:5215–7.Google Scholar

  • [113]

    Rohr TE, Cotton T, Fan N, Tarcha PJ. Immunoassay employing surface-enhanced Raman spectroscopy. Anal Biochem 1989;182:388–98.Google Scholar

  • [114]

    Nie S, Emory SR. Probing single molecules and single nanoparticles by surface-enhanced Raman scattering. Science 1997;275:1102–6.Google Scholar

  • [115]

    Qiu T, Zhang W, Chu PK. Recent progress in fabrication of anisotropic nanostructures for surface-enhanced Raman spectroscopy. Recent Pat Nanotechnol 2009;3:10–20.Google Scholar

  • [116]

    Le Ru EC, Etchegoin PG. EM enhancements and plasmon resonances: examples and discussions. In: Principles of Surface-Enhanced Raman Spectroscopy and related plasmonic effects. Great Britain, Elsevier, 2009.Google Scholar

  • [117]

    Benford M, Cote G, Kameoka J, Wang M. Raman detection in microchips and microchannels. In: Hawkins AR, Schmidt H, eds. Handbook of optofluidics. Oxon, CRC Press, 2010, 17-1–17-25.Google Scholar

  • [118]

    Jackson JB, Westcott SL, Hirsch LR, West JL, Halas NJ. Controlling the surface enhanced Raman effect via the nanoshell geometry. Appl Phys Lett 2003;82:257–9.Google Scholar

  • [119]

    Huebner U, Schneidewind H, Cialla D, et al. Fabrication of regular patterned SERS arrays by electron beam lithography. Proc SPIE 2010;7715:1–7.Google Scholar

  • [120]

    Bhandari D, Wells SM, Polemi A, Kravchenko II, Shuford KL, Sepaniak MJ. Stamping plasmonic nanoarrays on SERS-supporting platforms. J Raman Spectrosc 2011;42:1916–24.Google Scholar

  • [121]

    Krishnamoorthy S, Krishnan S, Thoniyot P, Low HY. Inherently reproducible fabrication of plasmonic nanoparticle arrays for SERS by combining nanoimprint and copolymer lithography. ACS Appl Mater Interfaces 2011;3:1033–40.Google Scholar

  • [122]

    Chattopadhyay S, Lo H-C, Hsu C-H, Chen L-C, Chen K-H. Surface-enhanced Raman spectroscopy using self-assembled silver nanoparticles on silicon nanotips. Chem Mater 2005;17:553–9.Google Scholar

  • [123]

    Fang J, Du S, Lebedkin S, et al. Gold mesostructures with tailored surface topography and their self-assembly arrays for surface-enhanced Raman spectroscopy. Nano Lett 2010;10:5006–5013.Google Scholar

  • [124]

    Liu Y-J, Chu HY, Zhao Y-P. Silver nanorod array substrates fabricated by oblique angle deposition: morphological, optical, and SERS characterizations. J Phys Chem C 2010;8176–83.Google Scholar

  • [125]

    Fan J-G, Zhao Y-P. Gold-coated nanorod arrays as highly sensitive substrates for surface-enhanced Raman spectroscopy. Langmuir 2008;24:14172–5.Google Scholar

  • [126]

    Cialla D, Hübner U, Schneidewind H, Möller R, Popp J. Probing innovative microfabricated substrates for their reproducible SERS activity. Chem Phys Chem 2008;9:758–62.Google Scholar

  • [127]

    You H, Ji Y, Wang L, et al. Interface synthesis of gold mesocrystals with highly roughened surfaces for surface-enhanced Raman spectroscopy. J Mater Chem 2012;22:1998–2006.Google Scholar

  • [128]

    Hüttner W, Christou K, Göhmann A, Beushausen V, Wackerbarth H. Implementation of substrates for surface-enhanced Raman spectroscopy for continuous analysis in an optofluidic device. Microfluidics Nanofluidics 2012;12:521–7.Google Scholar

  • [129]

    Copp J-P, Xu Z, Chen Y, Liu GL. Metallic nanocone array photonic substrate for high-uniformity surface deposition and optical detection of small molecules. Nanotechnology 2011;22:1–7.Google Scholar

  • [130]

    Zhixun L, Yan F. SERS of Gold/C60(/C70) nano-clusters deposited on floppy disk and hard disk. Chem Phys 2006;321:86–90.Google Scholar

  • [131]

    Lin Y, Bunker CE, Fernando KAS, Connell JW. Aqueously dispersed silver nanoparticle-decorated boron nitiride nanosheets for reusable, thermal oxidation-resistant surface enhanced Raman spectroscopy (SERS) devices. ACS Appl Mater Interfaces 2012;4:1110–7.Google Scholar

  • [132]

    März A, Rösch P, Henkel T, Malsch D, Popp J. Lab-on-a-chip surface-enhanced Raman spectroscopy. In: Fritzsche W, Popp J, eds. Optical nano- and microsystems for bioanalytics. Vol. 10, Berlin Heidelberg, Springer, 2012, 229–45.Google Scholar

  • [133]

    Zhang JZ, Noguez C. Plasmonic optical properties and applications of metal nanostructures. Plasmonics 2008;127–50.Google Scholar

  • [134]

    Marimuthu A, Christopher P, Linic S. Design of plasmonic platforms for selective molecular sensing based on surface-enhanced Raman spectroscopy. J Phys Chem C 2012;116:9824–9.Google Scholar

  • [135]

    Cassar RN, Graham D, Larmour I, Wark AW, Faulds K. Synthesis of size tunable monodispersed silver nanoparticles and the effect of size on SERS enhancement. Vib Spectro 2014;71:41–6.Google Scholar

  • [136]

    Walton BM, Huang PJ, Kameoka J, Cote GL. Use of a micro- to nanochannel for the characterization of surface-enhanced Raman spectroscopy signals from unique functionalized nanoparticles. J Biomed Opt 2016;21:85006.Google Scholar

  • [137]

    Liu X, Atwater M, Wang J, Huo Q. Extinction coefficient of gold nanoparticles with different sizes and different capping ligands. Colloids Surf B Biointerfaces 2007;58:3–7.Google Scholar

  • [138]

    Zhang Q, Xie J, Yu Y, Lee JY. Monodispersity control in the synthesis of monometallic and bimetallic quasi-spherical gold and silver nanoparticles. Nanoscale 2010;2:1962–75.Google Scholar

  • [139]

    Wang YL, Schlucker S. Rational design and synthesis of SERS labels. Analyst 2013;138:2224–38.Google Scholar

  • [140]

    Willets KA, Van Duyne RP. Localized surface plasmon resonance spectroscopy and sensing. Annu Rev Phys Chem 2007;58:267–97.Google Scholar

  • [141]

    Storhoff JJ, Lazarides AA, Mucic RC, Mirkin CA, Letsinger RL, Schatz GC. What controls the optical properties of DNA-linked gold nanoparticle assemblies? J Am Chem Soc 2000;122:4640–50.Google Scholar

  • [142]

    Mirkin CA, Letsinger RL, Mucic RC, Storhoff JJ. A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 1996;382:607–9.Google Scholar

  • [143]

    Cao YC, Jin R, Mirkin CA. Nanoparticles with Raman spectroscopic fingerprints for DNA and RNA detection. Science 2002;297:1536–40.Google Scholar

  • [144]

    Wang J, Lin D, Lin J, et al. Label-free detection of serum proteins using surface-enhanced Raman spectroscopy for colorectal cancer screening. J Biomed Optics 2014;19:087003.Google Scholar

  • [145]

    Rygula A, Majzner K, Marzec KM, Kaczor A, Pilarczyk M, Baranska M. Raman spectroscopy of proteins: a review. J Raman Spectro 2013;44:1061–76.Google Scholar

  • [146]

    Xiao SJ, Wieland M, Brunner S. Surface reactions of 4-aminothiophenol with heterobifunctional crosslinkers bearin both succinimidl ester and maleimide for biomolecular immobilization. J Colloid Interface Sci 2005;290:172–83.Google Scholar

  • [147]

    Chou IH, Benford M, Beier HT, et al. Nanofluidic biosensing for ß-amyloid detection using surface enhanced raman spectroscopy. Nano Lett 2008;8:1729–35.Google Scholar

  • [148]

    Feng S, Wang W, Tai IT, Chen G, Chen R, Zeng H. Label-free surface-enhanced Raman spectroscopy for detection of colorectal cancer and precursor lesions using blood plasma. Biomed Optics Express 2015;6:3494–502.Google Scholar

  • [149]

    Vo-Dinh T. Nanobiosensing using plasmonic nanoprobes. IEEE J Sel Top Quantum Electron 2008;14:198–205.Google Scholar

  • [150]

    Marks HL, Pishko MV, Jackson GW, Coté GL. Rational design of a bisphenol a aptamer selective surface-enhanced raman scattering nanoprobe. Anal Chem 2014;86:11614–9.Google Scholar

  • [151]

    Marks H, Mabbott S, Jackson GW, Graham D, Cote GL. SERS active colloidal nanoparticles for the detection of small blood biomarkers using aptamers. Proc. SPIE 9338, Colloidal Nanoparticles for Biomedical Applications 2015;9338:93381C–93381C-5.Google Scholar

  • [152]

    Schutz M, Kustner B, Bauer M, Schmuck C, Schlucker S. Synthesis of glass-coated SERS nanoparticle probes via SAMs with terminal SiO2 precursors. Small 2010;6:733–7.Google Scholar

  • [153]

    Kustner B, Gellner M, Schutz M, et al. SERS labels for red laser excitation: silica-encapsulated SAMs on tunable gold/silver nanoshells. Angew Chem Int Ed Engl 2009;48:1950–3.Google Scholar

  • [154]

    Schlucker S. SERS microscopy: nanoparticle probes and biomedical applications. Chemphyschem 2009;10:1344–54.Google Scholar

  • [155]

    Jehn C, Kustner B, Adam P, et al. Water soluble SERS labels comprising a SAM with dual spacers for controlled bioconjugation. Phys Chem Chem Phys 2009;11:7499–504.Google Scholar

  • [156]

    Donnelly T, Faulds K, Graham D. Investigation of silver nanoparticle assembly following hybridization with different lengths of DNA. Part Part Syst Charact 2016;33:404–11.Google Scholar

  • [157]

    Mabbott S, Thompson D, Sirimuthu N, McNay G, Faulds K, Graham D. From synthetic DNA to PCR product: detection of fungal infections using SERS. Faraday Discuss 2016;187:461–72.Google Scholar

  • [158]

    Simpson J, Craig D, Faulds K, Graham D. Mixed-monolayer glyconanoparticles for the detection of cholera toxin by surface enhanced Raman spectroscopy. Nanoscale Horizons 2016;1:60–3.Google Scholar

  • [159]

    Gracie K, Lindsay D, Graham D, Faulds K. Bacterial meningitis pathogens identified in clinical samples using a SERSDNA detection assay. Anal Methods 2015;7:1269–72.Google Scholar

  • [160]

    Chung E, Jeon J, Yu J, Lee C, Choo J. Surface-enhanced Raman scattering aptasensor for ultrasensitive trace analysis of bisphenol A. Biosens Bioelectron 2015;64:560–5.Google Scholar

  • [161]

    Mahmoudi M, Sant S, Wang B, Laurent S, Sen T. Superparamagnetic iron oxide nanoparticles (SPIONs): development, surface modification and applications in chemotherapy. Adv Drug Deliv Rev 2011;63:24–46.Google Scholar

  • [162]

    Büchner T, Drescher D, Merk V, et al. Biomolecular environment, quantification, and intracellular interaction of multifunctional magnetic SERS nanoprobes. Analyst 2016;141:5096–106.Google Scholar

  • [163]

    Ramadan Q, Gijs MAM. Simultaneous sample washing and concentration using a “trapping-and- releasing” mechanism of magnetic beads on a microfluidic chip. Analyst 2011;136:1157–66.Google Scholar

  • [164]

    Kumar GVP, Rangarajan N, Sonia B, Deepika P, Rohman N, Narayana C. Metal-coated magnetic nanoparticles for surface enhanced Raman scattering studies. Bull Mater Sci 2011;34:207–16.Google Scholar

  • [165]

    Wang C, Li P, Wang J, et al. Polyethylenimine-interlayered core-shell-satellite 3D magnetic microspheres as versatile SERS substrates. Nanoscale 2015;7:18694–707.Google Scholar

  • [166]

    Zhang X, Zhu Y, Yang X, Zhou Y, Yao Y, Li C. Multifunctional Fe3O4@TiO2@Au magnetic microspheres as recyclable substrates for surface-enhanced Raman Scattering. Nanoscale 2014;6:5971–9.Google Scholar

  • [167]

    Sun LJ, He J, An SS, Zhang JW, Ren D. Facile one-step synthesis of Ag@Fe3O4 core-shell nanospheres for reproducible SERS substrates. J Mol Struct 2013;1046:74–81.Google Scholar

  • [168]

    Wang J, Wu X, Wang C, et al. Magnetically assisted surface-enhanced Raman spectroscopy for the detection of Staphylococcus aureus based on aptamer recognition. ACS Appl Mater Interfaces 2015;7:20919–29.Google Scholar

  • [169]

    Ge M, Wei C, Xu M, et al. Ultra-sensitive magnetic immunoassay of HE4 based on surface enhanced Raman spectroscopy. Anal Methods 2015;7:6489–95.Google Scholar

  • [170]

    He J, Li G, Hu Y. Aptamer recognition induced target-bridged strategy for proteins detection based on magnetic chitosan and silver/chitosan nanoparticles using surface-enhanced Raman spectroscopy. Anal Chem 2015;87:11039–47.Google Scholar

  • [171]

    Guarrotxena N, Liu B, Fabris L, Bazan GC. Antitags: nanostructured tools for developing SERS-based ELISA analogs. Adv Mater 2010;22:4954–8.Google Scholar

  • [172]

    Bhardwaj V, Srinivasan S, McGoron AJ. On-chip surface-enhanced Raman spectroscopy (SERS)-linked immuno-sensor assay (SLISA) for rapid environmental-surveillance of chemical toxins. Adv Environ Chem Biol Sens Technol Xii, 2015;9486.Google Scholar

  • [173]

    Chon H, Wang R, Lee S, et al. Clinical validation of surface-enhanced Raman scattering-based immunoassays in the early diagnosis of rheumatoid arthritis. Anal Bioanal Chem 2015;407:8353–62.Google Scholar

  • [174]

    Heydari E, Thompson D, Graham D, Cooper JM, Clark AW. An engineered nano-plasmonic biosensing surface for colorimetric and SERS detection of DNA-hybridization events. Proc. SPIE 9340, Plasmonics in Biology and Medicine XII. 2015; 9340:93400O–93400O-6.Google Scholar

  • [175]

    Krpetic Ž, Guerrini L, Larmour IA, Reglinski J, Faulds K, Graham D. Importance of nanoparticle size in colorimetric and SERS-based multimodal trace detection of Ni(II) ions with functional gold nanoparticles. Small 2012;8:707–14.Google Scholar

  • [176]

    Jang H, Hwang EY, Kim Y, Choo J, Jeong J, Lim DW. Surface-enhanced Raman scattering and fluorescence-based dual nanoprobes for multiplexed detection of bacterial pathogens. J Biomed Nanotechnol 2016;12:1938–51.Google Scholar

  • [177]

    Whitesides GM. The origins and the future of microfluidics. Nature 2006;442:368–73.Google Scholar

  • [178]

    Chen L, Choo J. Recent advances in surface-enhanced Raman scattering detection technology for microfluidic chips. Electrophoresis 2008;29:1815–28.Google Scholar

  • [179]

    Zhou Q, Kim T. Review of microfluidic approaches for surface-enhanced Raman scattering. Sens Actuators B Chem 2016;227:504–14.Google Scholar

  • [180]

    Lim C, Hong J, Chung BG, deMello AJ, Choo J. Optofluidic platforms based on surface-enhanced Raman scattering. Analyst 2010;135:837–44.Google Scholar

  • [181]

    Chon H, Lim C, Ha S-M, et al. On-chip immunoassay using surface-enhanced Raman scattering of hollow gold nanospheres. Anal Chem 2010;82:5290–5.Google Scholar

  • [182]

    Wilson R, Bowden SA, Parnell J, Cooper JM. Signal enhancement of surface enhanced Raman scattering and surface enhanced resonance raman scattering using in situ colloidal synthesis in microfluidics. Anal Chem 2010;82:2119–23.Google Scholar

  • [183]

    Quang LX, Lim C, Seong GH, Choo J, Do KJ, Yoo S-K. A portable surface-enhanced Raman scattering sensor integrated with a lab-on-a-chip for field analysis. Lab Chip 2008;8:2214–9.Google Scholar

  • [184]

    Wang M, Jing N, Chou IH, Cote GL, Kameoka J. An optofluidic device for surface enhanced Raman spectroscopy. Lab Chip 2007;7:630–2.Google Scholar

  • [185]

    Yazdi SH, White IM. A nanoporous optofluidic microsystem for highly sensitive and repeatable surface enhanced Raman spectroscopy detection. Biomicrofluidics 2012;6:0104105.Google Scholar

  • [186]

    Gao R, Ko J, Cha K, et al. Fast and sensitive detection of an anthrax biomarker using SERS-based solenoid microfluidic sensor. Biosens Bioelectron 2015;72:230–6.Google Scholar

  • [187]

    Liu GL, Lee LP. Nanowell surface enhanced Raman scattering arrays fabricated by soft-lithography for label-free biomolecular detections in integrated microfluidics. Appl Phys Lett 2005;87:074101.Google Scholar

  • [188]

    Oh YJ, Jeong KH. Optofluidic SERS chip with plasmonic nanoprobes self-aligned along microfluidic channels. Lab Chip 2014;14:865–8.Google Scholar

  • [189]

    Li X, Ballerini DR, Shen W. A perspective on paper-based microfluidics: current status and future trends. Biomicrofluidics 2012;6:011301.Google Scholar

  • [190]

    Dungchai W, Chailapakul O, Henry CS. A low-cost, simple, and rapid fabrication method for paper-based microfluidics using wax screen-printing. Analyst 2011;136:77–82.Google Scholar

  • [191]

    Li X, Tian J, Garnier G, Shen W. Fabrication of paper-based microfluidic sensors by printing. Colloids Surf B Biointerfaces 2010;76564–70.Google Scholar

  • [192]

    Martinez AW, Phillips ST, Whitesides GM, Carrilho E. Diagnostics for the developing world: microfluidic paper-based analytical devices. Anal Chem 2010;82:3–10.Google Scholar

  • [193]

    Yu WW, White IM. A simple filter-based approach to surface enhanced Raman spectroscopy for trace chemical detection. Analyst 2012;137:1168–73.Google Scholar

  • [194]

    Chen J, Wu X, Huang Y-W, Zhao Y. Detection of E. coli using SERS active filters with silver nanorod array. Sens Actuators B Chem 2014;191:485–90.Google Scholar

  • [195]

    Gao S, Glasser J, He L. A filter-based surface enhanced Raman spectroscopic assay for rapid detection of chemical contaminants. J Vis Experiments 2016;108:e53791.Google Scholar

  • [196]

    Zheng G, Polavarapu L, Liz-Marzan LM, Pastoriza-Santos I, Perez-Juste J. Gold nanoparticle-loaded filter paper: a recyclable dip-catalyst for real-time reaction monitoring by surface enhanced Raman scattering. Chem Commun 2015;51:4572–5.Google Scholar

  • [197]

    Cheng M-L, Tsai B-C, Yang J. Silver nanoparticle-treated filter paper as a highly sensitive surface-enhanced Raman scattering (SERS) substrate for detection of tyrosine in aqueous solution. Anal Chim Acta 2011;708:89–96.Google Scholar

  • [198]

    Liu Q, Wang J, Wang B, et al. Paper-based plasmonic platform for sensitive, noninvasive, and rapid cancer screening. Biosens Bioelectron 2014;54:128–34.Google Scholar

  • [199]

    Ngo YH, Then WL, Shen W, Garnier G. Gold nanoparticles paper as a SERS bio-diagnostic platform. J Colloid Interface Sci 2013;409:59–65.Google Scholar

  • [200]

    Yu WW, White IM. Inkjet printed surface enhanced Raman spectroscopy array on cellulose paper. Anal Chem 2010;82:9626–30.Google Scholar

  • [201]

    Yu WW, White IM. Inkjet-printed paper-based SERS dipsticks and swabs for trace chemical detection. Analyst 2013;138:1020–5.Google Scholar

  • [202]

    Torul H, Çiftçi H, Çetin D, Suludere Z, Boyaci IH, Tamer U. Paper membrane-based SERS platform for the determination of glucose in blood samples. Anal Bioanal Chem 2015;407:8243–251.Google Scholar

  • [203]

    Qu L-L, Li D-W, Xue J-Q, Zhai W-L, Fossey JS, Long Y-T. Batch fabrication of disposable screen printed SERS arrays. Lab Chip 2012;12:876–81.Google Scholar

  • [204]

    Chin CD, Linder V, Sia SK. Commercialization of microfluidic point-of-care diagnostic devices. Lab Chip 2012;12:2118–34.Google Scholar

  • [205]

    O’Farrell B. Evolution in lateral flow–based immunoassay systems. In: Wong R, Tse H, eds. Lateral flow immunoassay. New York, Springer, 2009, 1–33.Google Scholar

  • [206]

    Hwang J, Lee S, Choo J. Application of a SERS-based lateral flow immunoassay strip for the rapid and sensitive detection of staphylococcal enterotoxin B. Nanoscale 2016;8:11418–25.Google Scholar

  • [207]

    Fu X, Cheng Z, Yu J, Choo P, Chen L, Choo J. A SERS-based lateral flow assay biosensor for highly sensitive detection of HIV-1 DNA. Biosens Bioelectron 2016;78:530–7.Google Scholar

  • [208]

    Choi S, Hwang J, Lee S, Lim DW, Joo H, Choo J. Quantitative analysis of thyroid-stimulating hormone (TSH) using SERS-based lateral flow immunoassay. Sens Actuators B Chem 2017;240:358–64.Google Scholar

Back to top