Functional Nucleic-Acid-Based Sensors for Environmental Monitoring

Functional Nucleic-Acid-Based Sensors for EnvironmentalMonitoring

Arghya Sett & Suradip Das & Utpal Bora

Received: 30 January 2014 /Accepted: 19 May 2014# Springer Science+Business Media New York 2014

Abstract Efforts to replace conventional chromatographic methods for environmental mon-itoring with cheaper and easy to use biosensors for precise detection and estimation ofhazardous environmental toxicants, water or air borne pathogens as well as various otherchemicals and biologics are gaining momentum. Out of the various types of biosensorsclassified according to their bio-recognition principle, nucleic-acid-based sensors have shownhigh potential in terms of cost, sensitivity, and specificity. The discovery of catalytic activitiesof RNA (ribozymes) and DNA (DNAzymes) which could be triggered by divalent metallicions paved the way for their extensive use in detection of heavy metal contaminants inenvironment. This was followed with the invention of small oligonucleotide sequences calledaptamers which can fold into specific 3D conformation under suitable conditions after bindingto target molecules. Due to their high affinity, specificity, reusability, stability, and non-immunogenicity to vast array of targets like small and macromolecules from organic, inor-ganic, and biological origin, they can often be exploited as sensors in industrial wastemanagement, pollution control, and environmental toxicology. Further, rational combinationof the catalytic activity of DNAzymes and RNAzymes along with the sequence-specificbinding ability of aptamers have given rise to the most advanced form of functional nucleic-acid-based sensors called aptazymes. Functional nucleic-acid-based sensors (FNASs) can beconjugated with fluorescent molecules, metallic nanoparticles, or quantum dots to aid in rapiddetection of a variety of target molecules by target-induced structure switch (TISS) mode.Although intensive research is being carried out for further improvements of FNAs as sensors,challenges remain in integrating such bio-recognition element with advanced transductionplatform to enable its use as a networked analytical system for tailor made analysis ofenvironmental monitoring.

Appl Biochem BiotechnolDOI 10.1007/s12010-014-0990-3

All authors contributed equally.

A. Sett : S. Das :U. Bora (*)Bioengineering Research Laboratory, Department of Biotechnology, Indian Institute of TechnologyGuwahati, Guwahati, Assam 781039, Indiae-mail: [email protected]

Utpal Borae-mail: [email protected]

S. Das : U. BoraMugagen Laboratories Pvt. Ltd., Technology Incubation Centre, Indian Institute of Technology Guwahati,Guwahati, Assam 781039, India

Keywords Functional nucleic acids . Sensors . Environmental monitoring

Introduction

Sensors are analytical tools that react to any physical, chemical, or biological stimuli andgenerate detectable signals [1]. They are a critical extension of human perception and sensationin many aspects of the life. This is largely because human sensory organs are much lesssensitive to the chemical or biological environment than to the physical environment (e.g.,light, pressure, temperature, and humidity). Human eye is the most important sensory organ.While focal length of human eye is 3.2 mm and aperture is f2.1–f8.3, it is magnified by variousmicroscopies like compound microscopy and cutting-edge electron microscopy (SEM, TEM,STM). Alike, there is persistent development in sensor technology; new-age sophisticated,sensitive biosensors are based on functional nucleic acid (FNA) dependent platform. There is aplenty of essential chemical or biological compositions which are core, basic, and indispensableelements of the quality of life. The symbiotic relationship between human and its niche leadsto consequences into man-made perturbations to environmental systems. So, early detection ofemerging environmental pollutants, contaminants, and toxins in natural or artificial environ-ments is important to identify and quantify exposure risks and to select remedial measure-ments. In an effort to identify and detect these environmental hazards, the ideal sensingtechnology should be capable to react to and diagnose the type of exposure within seconds.It can cover a wide spectrum of toxic agents, exhibits high sensitivity and reproducibility,requires minimum or no sample preparation, consumes minimal amounts of reagents, and iseasy and portable for field operations [2]. The environmental analytical communities alwaysquest for portable, reusable analytical devices which can offer reliable, precise, on-site analysisfor a variety of matrices and a host of diverse analytes. Rapid progress in nucleic acidchemistry, in-depth analysis of basic elements of life widened the range of biological recognitioncandidates and developments in microelectronics. Fiber optics technology optics technology hasalso expanded the capability of signal-transduction platform. Current trends of chemical detectionand quantification strategies like high-resolution gas chromatography, high-resolution massspectrometryanalysis [3], which are off-line and also often time consuming. Hence, to attainthe essential resolution, it does not permit for sufficiently rapid feedback to institute controls ornotify mitigation strategies. To encounter the current measurement challenges, cutting-edgesensors are currently designed and developed using novel biological element-based biosensors,autonomous cell-based toxicity monitoring, “lab-on-a-chip” devices rather than conventional/traditional techniques. Different variability like short lifetime, lack of genetic stability, strongbackground noise, and strong matrix effect on signal serves as a hindrance to chemicalinterpretation of traditional biosensors. Hence, in recent era, cell-free system-based biosensorshave been emerging to overcome the limitations related to whole cell systems.

A few such type of cell-free systems are embodied by single-stranded oligonucleotidescomposed of DNAzymes and RNAzymes that bind to a variety of molecules with precisespecificity [4]. Although first reported in late 1990s, they have been exploited recently asligands in biosensors with applications in the areas of diagnostics and therapeutics owing totheir high selectivity, specificity, high reproducibility, cost-effectiveness, and ease of use.Although nucleic acid oligomers hold less complexity and chemical functionalities comparedto proteins, binding affinity toward the target molecule of the functional nucleic acids (FNAs)can also rival that of antibody which is produced in-vivo. There are certain drawbacks ofnucleic acids as compared to amino acids which have greater torsional freedom and confor-mational flexibility. Modification with additional chemical groups or internal structure

Appl Biochem Biotechnol

dynamics escalates the conformational freedom and binding capacity of FNAs with theirtargets [5]. Moreover, diverse chemical groups can be varied to obtain a better and suitableFNA-target contact in complexes of traditionally known nucleoside structure. Merely, a smallmodification of nucleosides greatly increases the complexity of a library. Altered structure ofmodified nucleosides offer greater conformational diversity of FNAs and thereby increase thepotential of a library. Modified FNA structures can have more stable conformation, whichmake the FNAs like DNAzyme and RNAzyme more custom-made. Alteration and modifica-tions in the internal structures of DNAzyme and RNAzyme can also increase the affinity for atarget. Lastly, modified FNAs can be more resistant to hydrolysis, nuclease resistant, andthermally stable which expands their applicability in the field of sensor technology. Thedurable, sensitive, and economical FNA-based sensors have now emerged as a novel combattool against the ever-increasing environmental pollutants such as toxic heavy metals, airborneand waterborne microbes, and other biological toxins in this modern era of industrialization.

Types of FNAs

Discovery of ribozymes by Cech and Altman altered the concept that all biocatalysts areprotein molecules [6]. Earlier nucleic acids were known as hereditary material for storage ofgenetic information of life. Functional nucleic acids are not only “blueprint of life” but can alsoact as catalytic and ligand binding element. FNAs are broadly classified in two categories: (a)catalytic FNAs (DNAzyme and RNAzyme) and (b) non-catalytic FNAs (aptamers andspiegelmers) (Fig. 1).

Fig. 1 Types of various functional nucleic acids (FNAs)


The catalytic FNAs act in two successive steps: first, recognition of the sequence and thencleavage of the sequence at a specific site. Naturally found RNAzymes can catalyze RNAsplicing, cleavage, and peptide bond formation in ribosome machinery [7–9], but artificiallyderived RNAzymes can catalyze a vast array of reactions such as RNA cleavage, ligation,polymerization, alcohol dehydrogenation, porphyrinmetalation, acyl transfer and degradationof amide, urea, and other small macromolecules [10–17]. Although DNAzymes, lacking 2′hydroxyl group, are comparatively new, they also can catalyze a variety of reactions rangingfrom ligation and cleavage reactions of both RNA and DNA, RNA branching and lariat RNAformation, phosphorylation, deglycolation, and other enzyme reaction as of peroxidases. Thetwo most comprehensively well studied RNA-cleaving DNAzymes are 8–17 and 10–23 tilldate and have the ability to carry out sequence-specific cleavage of all RNA and chimericRNA/DNA substrates. The complimentary sequence specific against target lies between 8–17and 10–23 bp catalytic core of deoxyribozyme. The substrate target for each DNAzyme is achimeric DNA/RNA molecule containing a ribonucleotide (rA/rT/rG/rC) as the cleavage site.These man-made ribozymes, deoxyribozymes, are all key players along with naturally found“molecular wonders” riboswitches.

Non-catalytic FNAs like aptamer are also widely exploited in sensor technology. DNA orRNA aptamers can be custom-made, tagged with different fluorophores, quenchers, or metallicnanoparticles for precise and sensitive detection of environmental pollutants. Suitable surfaceimmobilization strategy provides specific adsorption of nucleic acids and also decreases thebackground noise. Among the physical adsorption and chemical-binding strategies [18], theformation of a gold–thiol bond is comparatively easy and stable and is the most popular for theimmobilization of the DNA strand. The carbodiimide bond formed between –COOH and –NH2 is also used to introduce probe DNA. Besides, the biotinylated single-stranded DNA(ssDNA) aptamer can be immobilized on a streptavidin modified electrode through biotin-avidin interaction [19].

However, the application of aptamers as sensing molecule is hindered as a result of theirsusceptibility to nuclease degradation. The incorporation of 2′-modified RNA libraries ormodified base, sugar or phosphate-backbone improves the biological half-life significantly.Introduction of chiral principles into the in vitro selection process aids to identify nucleaseresistant oligonucleotides. Among few applications, one of the approaches has led to theidentification of L-enantiomeric RNA or DNA ligands termed “spiegelmers” that bind toarginine [20], adenosine [21], and vasopressin [22]. Spiegelmers can identify their targets inthe same way as aptamers, but they are resistant to enzymatic degradation by nucleases. Non-catalytic FNAs can recognize a wide range of target molecules from biological origin to non-living molecules. To gain advantage of high sensitivity of catalytic FNAs, aptamers have nowbeen coupled with DNAzymes, RNAzymes, and other catalytic modules to broaden itsapplication as a sensing tool.

In Vitro Selection of FNA from Combinatorial Library

Rational methods can be applied to reformulate existing natural ribozymes or to createribozymes with completely different catalytic and kinetic features [23, 25]. The simpleststrategy is composed of splitting a contiguous polynucleotide chain to create a fragmentedribozyme. The fragments can then be reassembled to form an active multicomponent catalyticribozyme. Simple secondary structure elements can be created by following the Watson-Crickbase complementation. These strategies are used routinely to create ribozymes that offermultiple turnover kinetics or altered substrate specificity. However, as with the rational design


of ribozymes, control over the precise positioning of RNA functional moiety in three-dimensional structures remains beyond the reach of current rational design techniques [24,26]. An alternative or advancement to rational design is the onset of iterative selection methodsthat isolate catalytic molecules from mutagenized or randomized sequence pools of RNA orDNA. This in vitro selection approach based on the probability that a given pool of randomsequence molecules will include individual elements that can perform the function of interest.As an instance, in case of the conserved catalytic core (13 nucleotides) of hammerhead self-cleaving 10–23 ribozyme, it is expected to occur with a frequency of 1 in every 67 millionrandom-sequence RNAs. Therefore, in a pool of 1015molecules, approximately 15 millionelements can possess the hammerhead catalytic core, and some of them can efficiently catalyzeRNA cleavage. Engineering new catalysts then has been reduced to the synthesis ofmutagenized ribozyme pools or random-sequence pools of nucleic acids.

But, there is always an uncertainty that a variety of catalytic molecules will be present in adiverse pool of nucleic acids. The more challenging issue is to screen or isolate those raremolecules containing the desired catalytic properties. This problem can be resolved if theelements that perform the desired catalysis can be separated from the remaining pool andsubsequently amplified using any of the methods for the replication of RNA and DNA. Thisprocess of selective amplification or “in vitro selection” can be used in an iterative fashion toscreen rare molecules from large random pools of RNA or DNA. If a significant number ofmutations are incorporated to obtain customized candidate during selective amplification, thenthe process is typically termed as “in vitro evolution” to reflect the similarities between thisprocess and Darwinian evolution.

For metal ion-dependent DNAzyme selection, the process is target-dependent. For aninstance, the initial DNA pool containing 40–60-nucleotide-long random regions flanked by2 constant regions is taken. In the middle of this DNA strand, a putative cleavage site isintroduced in terms of ribo-adenosine (rA), since a ribonucleotide is ~100,000-fold moresusceptible to hydrolytic cleavage than a deoxyribonucleotide.

To obtain DNAzymes with high metal ion affinity, the metal concentration is decreasedafter each round of selection [7, 27] (Fig. 2). To increase the substrate specificity, negativeselection steps can be introduced to eliminate sequences that are also active with competingmetal ions [28]. After the activity of the DNA pool reached a plateau, the DNAs were clonedand sequenced. After accurate truncation, modification, and rational design of substratebinding sequences, a trans-cleavage DNAzyme is constructed. The strand having rA acts likesubstrate, and the other strands act like the enzyme. This selection protocol enable nucleic acidenzyme (NAE)-based biosensors to be more tailor-made and more versatile pertaining tovarious applications [29].

In combinatorial chemistry, any processes including Systemic Evolution of Ligands byEXperimental enrichment (SELEX) involve three steps: synthesis of an aptamer library,selection of screening, and structural analysis of the resulting enzyme-target complex.Aptamers obtained at the end of the SELEX process, high affinity and specificity for a targetand their postselection modification can further enhance these properties. In vitro selectioninitiates with a “random pool” containing typically 1014–15 different DNA sequences (depend-ing on the no. of random nucleotide bases). The pool is designed by flanking a randomsequence with two constant regions that are primer binding sites. After the “random pool” issynthesized, it is incubated with the substrate to carry out the desired reaction in the presenceof a cofactor. Since in vitro selection normally only utilizes a ssDNA, such ssDNA is thengenerated from the PCR products, and they are incubated under the desired condition with thesubstrate to start the next round of selection. After iterative rounds of selection, separation, andamplification, the large initial pool of random sequences can be reduced into a small


population of sequences that are enriched with substrate-specific aptamer. About 10–15 roundsof cyclic selection process are required to select the enriched molecule against the candidate.At the end of the process, the selected population is cloned and sequenced. The resultingsequences are grouped into different families based on sequence similarity, and each family istested for activity.

To identify a spiegelmer, the target of choice has to be prepared in its mirror-imageconfiguration. A standard in vitro selection scheme is performed to isolate an aptamer (D-nucleic acid) that binds to this mirror-image target. After synthesizing the isolated aptamersequence in its respective L-enantiomeric form, the resulting spiegelmer binds the natural target(in the natural configuration) with comparable affinity [30].

The selection of functional nucleic acids is an execution of the concept of “survival of thefittest molecule,” allowing researchers to find new candidate molecules with desirable activ-ities in the presence of intended targets.

Role of “Heavy Metals” in Environmental Pollution

The term “heavy metal” is conventionally used to define elements with high density andrelative atomic mass above 20 which is toxic to the biological system at low concentrations.Although the term heavy metal has been widely used by chemists and environmentalresearchers for the last 60 years, such a term has never been defined by International Unionof Pure and Applied Chemistry (IUPAC). According to an IUPAC technical report by JohnDuffus, the term heavy metal is both meaningless and misleading since it has been usedindiscriminately to indicate that all metals and semimetals which fall under this category aretoxic and hence pollutants. The article further goes on to suggest that the Lewis acidclassification on terms of electro-negativity provides a more accurate chemical basis fortoxicity assessment of elements without any reference to ‘heaviness’ [31]. Klopman classifiedelements into two broad classes in terms of electro-negativity where metals with orbitalelectro-negativities above 1.45 where grouped as Class Awhile those having electronegativitybelow −1.88 fell under Class B [32]. The Class B category includes the Lewis acids of largesize and high polarizibility like Cu(I), Pd, Ag, Cd, Ir, Pt, Au, Hg, Ti, and Pb(II). Further metals

Fig. 2 . Selection, optimization, and reformulation of a Mg2 + -dependent RNA-cleaving DNAzymes.Arrowhead identifies the cleavage site of DNAzyme. The sequences in box are representative of the new hairpinstructures that are found in optimized individuals. Encircled nucleotides are conserved (Reprinted with permis-sion Symons [7] Copyright © 1997, American Chemical Society)


with properties intermediary to Class A and B were classified as Borderline metals [33].Borderline metals included V, Cr, Mn, Fe(II), Co, Ni, Cu(II), Zn, Rh, Pb(IV), and Sn whichhave been reported to be toxic to biological systems above a threshold concentration. Arational explanation for biological toxicity of certain metals was provided by Nieboer andRichadson. It was hypothesized that Borderline and Class B ions similar in size to Ca (II) cancause structural damage to cell membranes due to their affinity for phosphate groups and non-oxygen carriers present in the membrane [34].

Escalating industrial activities like mining, corrosion, fossil fuel combustion, agriculture,and sludge dumping with increased metallurgical explorations have contributed to the accu-mulation of Class B and Borderline metals in our environment well and above safety levels.These metals tend to accumulate in the environment silently under our eyes unlike otherenvironmental pollutants like petroleum hydrocarbons or domestic and municipal waste wheredumping in the environment is noticeable. Extensive research on environmental pollutants andincreased awareness among people has helped to control the levels of such toxic metals. In aconcomitant fashion, the increased technological know-how has augmented the developmentof highly sensitive biosensors for detection and quantification of these toxic metals in ourenvironment at a very nascent stage.

Functional nucleic acids like DNAzymes and RNAzymes whose catalytic activity can beregulated by metals has been widely used as sensors in laboratory scale to detect toxic metalsin our environmental samples with high specificity and sensitivity. Further, the fact that metalscan induce conformational changes in aptamers has been exploited to develop aptasensors fordetection of toxic elements even in very low concentrations.

FNA-Based Sensors for Detection of Lead (Pb2+)

The major source of lead in our atmosphere is exhaust emission from automobiles running onleaded petrol. Such airborne lead contaminates our crop and reaches toxic levels due to bio-magnification along the food chain. Some of the other sources of lead in our environment arecontaminated water flowing through lead plumbing, lead containing paint and dust, and soilnear lead smelters [35]. Elevated levels of lead in the blood (>10 μg/dl in adults and >5 μg/dl inchildren) can lead to acute neuropathy and renal disorders. Lead interferes with many enzymeslike delta-aminolevulinic acid dehydratase, or ALAD, ferrochelataseetc which help in biosyn-thesis of hemoglobin by binding with the sulphahydryl groups present in these enzymes. Sincein many cases lead toxicity remains asymptomatic till almost 50 μg/dl, it is imperative to detectlead in blood and environments samples in trace amount at a nascent stage.

Almost a decade ago, a team from University of Illinois, USA, published a series of articlesdemonstrating the fabrication of DNAzyme based sensors for highly sensitive and specificdetection of Pb2+ ions from environmental samples. They showed that catalytic nucleic acidslike DNAzymes can be conjugated with gold nanoparticles to form colorimetric biosensor. Thenanoparticles were attached to 12mer-DNAmolecule having sequence complementary to 17DSregion of a 8–17 DNAzyme [36]. Optimization of reaction conditions and improved design ofthe DNAzyme enabled highly sensitive detection of Pb2+ (around 100 nM) within 10 min atambient temperature [37, 38]. Later, another group was able to achieve higher sensitivity (1 nM)of lead detection by immobilizing a thiol conjugated catalytic molecular beacon onto a goldsurface. Such fluorescent-based colorimetric biosensor could be regenerated to enable multiplecycles of sample testing. This was the first step toward the development of a stand-alonebiosensor for in situ detection of Pb [39]. The work was carried forward by Chang et al., whodeveloped a miniaturized lead sensor by integrating a lead specific DNAzyme to a nanocapillaryconnected microfluidic device to deliver small volume sample into a spatially confined detection


window [40]. The enhanced sensitivity, portability, and reusability of biosensors havingimmobilized DNAzymes encouraged further exploration of immobilization platforms.DNAzymes were immobilized on gold nanocapillary membranes [41], PMMA [42],polythiophene [43], and gold electrode surface for fabrication of DNA-Au bio barcodes [44].The barcode is formed by short oligonucleotides attached to 13-nm gold nanoparticles whichhybridize with the substrate strand of DNAzyme. Such electrochemical biosensor can detect verylow concentrations of lead (1 nM) in ground and drinking water. In an attempt to developportable biosensors capable of detecting lead in situ, Mazumdar et al. fabricated a simple dipstickbiosensor by immobilizing nanoparticle-DNAzyme conjugate on lateral flow devices for visualdetection of lead in paints at a minimum concentration of 5 μM within 10–15 min [45].

Signal amplification by quantitative PCR reaction has enabled development of label-freebiosensors for lead. Also, incorporation of quantitative PCR (QPCR) in the system resulted inhigh sensitivity (with a detection limit of 1 nM) of Pb2+ detection as well as good selectivityfor Pb2+ when challenged with coexisting ions [46]. In an alternative approach to developlabel-free biosensors, Guo et al. designed K+ stabilised G-quadruplex structures which boundto N-methyl mesoporphyrin IX (NMM) giving high fluorescence signal. In presence of Pb2+,lead could bind competitively to the G-quadruplex to form more compact DNA folds whichcould not bind to NMM leading to a decrease in fluorescence intensity. This allowedquantitative analysis of Pb2+ by simple “mix and detect” protocol with a detection limit of1 nM [47]. In a similar method, the dual input requirement of the G-quadruplex was utilized todesign a biosensor functioning as an AND gate. The biosensor was composed of a poly-G loopand a GR-5 DNAzyme stem which showed positive signal when Pb2+ and K + existsimultaneously in the system. The combination of hairpin DNA and GR-5 DNAzyme resultedin high selectivity of Pb2+ and a detection limit of 22.8 pM [48]. An electro-chemiluminescentlead biosensor based on GR-5 lead-dependent DNAzyme was reported to detect ultra lowconcentration of Pb2+ of almost 0.9 pM [49]. In a further innovation, similar G-quadruplexDNAzymes for lead detection were designed and integrated with multiple transducer plat-forms. Li et al. developed a colorimetric and chemiluminescent lead sensor using a commonG-quadruplex DNAzyme named PS2.M. The colorimetric system was found to have adetection limit of 32 nM whereas the chemiluminescent platform gave a better detection limitof 1 nM [50]. Similarly, another group later developed sensitive surface plasmon resonance(SPR) and electrochemical sensing platform for Pb2+ detection using Pb2+ sensitive DNAzymeand a hemin-G Quadruplex nanostructure. The SPR-based system offered ultralow detectionlimit of 5 fM whereas the electrochemical sensor had a detection limit of 1 pM. All thedifferent sensing platforms provided high selectivity for Pb2+ ions when challenged withcoexisting ions [51]. In an attempt to amplify the signal from sensors detecting lead, Zhuanget al. relied on the free energy-driven DNA hybridization chain reaction. They successfullydeveloped a magneto controlled electronic switch to respond to ultra-low concentration ofPb2+ ions in samples (37 pM) utilizing a lead-specific DNAzyme and two ferrocene-labeledDNA hairpins [52]. In a similar approach using streptavidin-conjugated magnetic beads tocapture biotinylated GR-5 DNAzyme attached to fluorophore, quantitative analysis of Pb2+

ions was performed by flow cytometric method. Such detection systems do away withconventional cuvette-based measurements which fail to provide accurate data for cloudyenvironmental samples due to high background as most of the light is absorbed or scattered.A high sensitivity of around 0.6 nM and high selectivity for Pb2+ was reported [53].

DNAzyme-based sensors are almost always associated with an enzyme strand and substratestrand. The use of two different oligonucleotides in one sensor not only made the systemcomplicated to design but also problematic to operate at elevated temperatures. To overcomethis, a Chinese group recently designed a DNAzyme-based fluorescent sensor for detection of


Pb2+ where both the enzyme and substrate strand was combined into a single oligonucleotidechain. The resulting intramolecular duplex structure allows the fluorophore and quencherattached at two ends to remain in close proximity at resting state. In presence of Pb2+, thefluorophore is released leading to increased fluorescence. The sensor was reported to be stableat various temperatures and operate with high selectivity and sensitivity of around 3.1 nM [54].

In a seminal study by Smirnov et al., the group reported for the first time the role of leadions in sequence-specific folding of DNA. Till then, DNA quadruplex structures were knownto be stabilized by divalent cations like K, Na, or NH3+. However, they found that Pb2+ couldbind with high affinity and induce folding of thrombin binding aptamers (TBAs) at micromo-lar concentrations. The study encouraged further research with lead specific aptamers fordetection and control of genotoxic effects of lead in the environment [55]. Conjugating aquantum dot and gold nanoparticle to TBAwas also reported to be able to detect micromolarlevels of lead via optical detection through fluorescent resonant energy transfer [56]. Similardetection systems have been studied for detection of various other heavy metals.

Various strategies that evolved with time for detection of Pb2+ from environmental samplesare represented as a collage in Fig. 3. Initially, DNAzymes were engineered so that repeatedheating and annealing steps for detection are no longer required. Further, the development offluorescence labeled as well as label-free sensors along with unimolecular (single strand)DNAzyme greatly enhanced the detection speed and simplified on-site detection. Also, thediscovery that Pb2+ could bind and induce folding of guanine rich DNA into G-quadruplexstructures lead to the development of novel sensors. Using gold nanoparticle conjugatedelectrodes, DNAzymes anchored to magnetic beads and highly specific aptamers as sensingplatform have greatly enhanced the sensitivity by stretching the detection limit to femtomolarconcentrations of Pb2 + .

FNA Based Sensors for Detection of Mercury (Hg2+)

The first report of an epidemic due to exposure to mercury came fromMinamata Bay, Japan, in1956. The poisoning was due to the highly toxic methyl-mercury which came from the nearby

Fig. 3 A schematic representation of the development of various detection platforms for environmentalmonitoring of Lead. The references are given in superscript while the year denotes the year of publication


Chisso Corporation factory manufacturing large amounts of acetaldehyde using mercurysulfate as a catalyst. As a side reaction of the process, the toxic methyl mercury was beingproduced in small quantities and entered the surrounding waters through factory effluents, thuscontaminating all fishes in the bay [57]. In the next 4 decades, 2,265 people suffered fromcrippling neurological diseases out of which 1,784 died. Chisso Corporation extended finan-cial compensation to over 10,000 victims and continued to do so till 2010 [58]. A similaroutbreak was reported few years later in Iraq. In the winter of 1971, several people from ruralareas of Iraq got admitted with symptoms of mercury poisoning. They were all victims ofmethyl-mercury poisoning resulting from the extensive use of fungicides (methoxypropyl-mercury and phenylmercury compounds) in wheat and barley cultivation [59]. The molecularbasis of mercury toxicity is based on its ability to irreversibly bind to selenium-containingenzymes or selenozymes like thioredoxin reductase which are essential in controlling oxidativedamage in cells [60]. The USFDA and Environmental Protection Agency (EPA) have fixed themaximum allowable level of mercury in food and drinking water to 1 ppm and 1 ppb,respectively [61].

One of the earliest attempts at detecting mercury from environmental samples was made byan American scientist in 1971. He developed a membrane probe-spectral emission typedetection system which could detect as low as 0.4 ng of metallic mercury from water samples[62]. Although the apparatus was unique in those days, it has rapidly become out-dated in themodern era of smart, portable biosensors based on advanced materials like functional nucleicacids. Mercury was reported to induce the folding of a DNA strand in the form of a hairpinwhich enabled its detection by FRET as the fluorophore and the quencher attached at two endscame in close proximity [63]. An array of nucleic acid sensors for mercury were developedutilizing mercury’s strong binding affinity for thymine residues and detect ultralow concen-trations of Hg2+. The presence of several thymine residues at the catalytic core of theDNAzyme rendered it inactive. However, even trace amount of Hg2+ was sufficient to formthymine–Hg2+–thymine bonds activating the fluorophore attached DNAzyme beacon leadingto increased fluorescence. The system could detect nanomolar concentrations Hg2+ andprovide more than 105 times selectivity for Hg2+ over other ions [64]. In a similar study, adetection system composed of polythymine oligonucleotides (T33) and TOTO-3 was designedto detect Hg2+ in pond water and batteries. TOTO-3 is a cyanine dye which exhibits increasedfluorescence when bound to double-stranded DNA (dsDNA). The presence of Hg2+ inaqueous solutions was found to induce folding of T33 into a double-stranded structure whichcould bind to TOTO-3 thereby showing a 1,000-fold increase in fluorescence. The studydemonstrated a rapid (<15 min), label-free and cost-effective method for detection of Hg2+

[65]. A simple label-free system with SYBR Green I as a signal reporter molecule for dsDNAwas able to detect Hg2+-induced folding of DNA strand with high sensitivity (0.5–1.33 nM)and specificity [66, 67]. In a fundamentally different approach, Xue et al. devised a one-step,room temperature colorimetric detection system for mercury. They used two separate DNAprobes conjugated with gold nanoparticle and a third linker probe that recognized the other twoprobe sequences. The probes were found to form stable DNA duplexes only in presence ofHg2+ with a detection limit of 3 μM by naked eye [68]. Nanoparticles conjugated withmercury-specific DNA or polythymine oligonucleotides (T33) were subsequently exploredfor detection of mercury reporting sub-micromolar sensitivity and high specificity for Hg2+

[69–71]. Moreover, addition of Mn2+ was reported to enhance the sensitivity of such systemsto a detection limit of 10 nM. This was probably due to the ability of Mn2+ to bind to thephosphate backbone of DNA, thereby stabilizing the Hg2+-T33 complex [72]. DNAzymes withG-quadruplex structure and showing peroxidase-like activity upon binding to hemin has alsobeen used for development of sensitive, label-free biosensors against Hg2+. Li et al. for the first


time, reported the use of such biosensors for detection of Hg2+ in pond water samples and wereable to achieve high sensitivity of 50 nM and high selectivity [73]. In a further improvement ofthe system, the G-quadruplex DNAzymes having peroxidise-like activity was used for visualdetection of Hg2+ utlilizing the tetramethyl benzidine (TMB)-H2O2 system with high sensi-tivity (100 nM) and high selectivity [74].

In a recent study, mercury-specific DNA aptamers were immobilized onto gold surface tomonitor the presence of Hg2+ in aqueous solutions by plasmon enhanced vibrational spectros-copy. A schematic of the sensor design is presented in Fig. 4. Direct detection of mercury wasachieved up to a very low concentration of 3.7 nM with high selectivity [75].

FNA-Based Sensors for Detection of Copper (Cu2+)

Copper is an essential nutrient to the body but excess of it may lead to serious toxicity issueslike hematemesis, hypotension, jaundice, and severe damage to the liver and brain. Most of thecopper in the body remains bound to a blood protein called ceruloplasmin which is essentialfor iron metabolism. The toxicity is caused by the unbound free copper molecules whichgenerate reactive oxygen species leading to damage in protein, nucleic acids, and lipids [76,77]. Natural sources of copper in the environment are from volcanic eruptions, forest fires, andwindblown dust. However, the major cause of copper poisoning in the environment are relatedto anthropogenic activities like use of pesticides, insecticides, and fungicides along withcopper mining and major electrical and metal manufacturing projects. [78]. According to theEPA, the maximum allowable limit for copper has been fixed at 1.3 mg/l or 1.3 ppm [79].

Pioneering scientist Ronald Breaker from Yale first isolated catalytic DNAs ordeoxyribozymes while working at Scripps Research Institute. The finding that copper ionsin presence of ascorbate or alone could modulate the catalytic activity of the DNA moleculelater became the foundation on which several nucleic-acid-based sensors were developed fordetection of copper ions [80–82]. Based on Breaker’s design of DNAzymes, a separate groupdeveloped a “turn on” fluorescent sensor for Cu2+ by labeling both ends of the substrate strandwith a fluorophore and quencher while the enzyme was tagged with another quencher. Thesensor could detect Cu2+ concentrations as low as 35 nM within 2–4 min with high specificity[83]. Such Cu2+-specific DNAzymes were immobilized onto aldehyde-coated slides for highthroughput microarray detection of the metal ion [84]. Label-free systems having DNAzymescoupled with gold nanoparticles were also explored and found to have nanomolar sensitivity,high specificity, and was successful when used in a pilot scale with surrounding water samples[85, 86]. A highly sensitive Cu2+ sensor having detection limit of 60 pM was later reportedusing dynamic light scattering technique to monitor the increase in hydrodynamic diameter ofnaked gold nanoparticles upon adsorbing ssDNA molecules released from Cu2+-dependentcleavage of DNAzyme [87]. Further innovative approaches have also been explored pertainingto various signal transduction and sensor systems for effective detection of copper. A lateralflow dipstick sensor was designed and composed of Cu2+-dependent DNAzyme and goldnanoparticles for visual detection of Cu2+ ions. The system was reported to have ultralowdetection limit of 10 nM with high specificity [88]. DNAzymes having G-quadruplex structurehave been effectively used for detection of other metal ions like Pb2+, Hg2+, etc. A G-quadruplex structure that binds with protoporphyrin IX (PPIX) leading to increased fluores-cence output was utilized for label-free detection of Cu2+. In presence of Cu2+, the fluores-cence of PPIX was quenched that enabled detection of Cu2+ in sample with high sensitivity(3 nM) [89]. A separate group engineered a complex unimolecular probe consisting of threedomains—a DNA-cleaving DNAzyme, a HRP mimicking DNAzyme, and a substrate strand.The system was engineered in such a way that in presence of Cu2+, the HRP mimicking


domain opened up, and its activity could be recorded by colorimetric methods. TheDNAzyme-based sensor could detect Cu2+ in water samples as low as 1 μM [90].

The use of aptamers in bio-sensing has emerged as an exciting field of research due to itshigh sensitivity, specificity and cost-effectiveness. Electrochemical aptasensors were exploredfor detection of Cu2+ and exhibited very high sensitivity of 0.1 pM, a wider detection range

Fig. 4 A schematic of mercury trapping by DNA aptamers. a Hg2+ ions are trapped inside the aptamers formingN- Hg2+-N bonds and conformational change of DNA. b A photo of Lake Kasumigaura and subaqueousenvironment. c Structural and chemical changes in the aptamer upon binding of Hg2+ ions lead to themodification in the electromagnetic field helping in detection of Hg2+ ions (Adapted with permission Hoang[75], Copyright © 2013, Nature Publishing Group)


(0.1 nM–10 μM) and excellent specificity. The sensor consisted of gold nanoparticles thatprovided higher surface area for anchorage of large number of aptamers, a catalytic self-cleaving DNA molecule and a ferrocene-labeled DNA for electrochemical transduction [91].

FNA-Based Sensors Against Multiple Targets

Through extensive research and development in sensor design and fabrication, rapid, on-site,and sensitive sensing solutions are available today for various toxic metals like Pb2+, Hg2+,Cu2+, etc. Several of the above discussed metal ions coexist in the environment. Thus, sensorscapable of simultaneous detection of multiple metal ions in the environment are essential. Withthis aim, an unlabeled, DNA-based sensor platform was engineered to detect Pb2+, Hg2+, andAg+ simultaneously in water and human serum samples. The sensor was designed to operate ina manner as described in Fig. 5 using appropriate masking agents [92]. Using a similarconcept, an electrochemical DNAzyme-based sensor was engineered for simultaneous detec-tion of Pb2+ and Hg2+ composed of Pb2+-specific DNAzyme and a Hg2+-specific aptamers. G-rich DNA and cysteine were alternatively used to mask Pb2+ and Hg2+, respectively. Target-induced conformational changes were monitored by electrochemical impedance spectroscopy(EIS) with a detection limit of 1 and 0.1 pM for Hg2+ and Pb2+, respectively [93].

FNA-Based Sensors for Detection of Other Toxic Metals

Several other metals like arsenic, uranium, cadmium, etc, accumulate in the environment dueto natural and anthropogenic activities and biomagnifies upon entering the food chain leadingto severe toxic effects. Early detection of such metals is essential for monitoring and control-ling their levels.

Aptamers have been shown to detect arsenic and cadmium from environmental samples atrelatively low concentrations. Arsenic binding aptamers were screened after several rounds ofin vitro selection, isolated and patented by a US group [94]. A separate group later demon-strated the detection of arsenic through colorimetric methods using arsenic specific aptamers

Fig. 5 A schematic representing sensor design for simultaneous detection of multiple heavy metals viz Pb2+,Hg2+, and Ag + using appropriate masking agents (Adapted with permission Lin [92], Copyright © 2011,American Chemical Society)


which remained bound to a cationic polymer and dispersed gold nanoparticles. Arsenic wasable to competitively bind to aptamers which allowed the cationic polymers to conjugate withgold nanoparticles. This lead to aggregation of the nanoparticles which could be detectedspectrophotometrically up to a detection limit of 5.3 ppb of arsenic [95]. The same group alsodemonstrated the capabilities of such sensor when integrated with other transduction systems.Arsenic could be detected from 40 ppb by naked eye to 0.6 and 0.77 ppb by colorimetric andresonance scattering techniques [96].

The bioavailability of water soluble uranium dioxide through increasing nuclear activitiesposes a major threat since this radioactive element is highly carcinogenic. A fluorescentbiosensor composed of DNAzyme tagged with a quencher and a substrate strand tagged withfluorophore and quencher was engineered against uranyl ions (UO2

+). The catalytic beaconwas “turned on” in presence of UO2

+ and exhibited a detection limit of 11 ppt with millionfoldsensitivity [97]. Recently, Tang et al. designed a fast, real-time electrochemical sensor foruranium based on ferrocene-labeled DNAzymes with a detection limit in nanomoles [98]. In anovel approach, another group developed a fluorophore tagged DNAzyme conjugated withgold nanoparticle for intracellular uranyl ion detection. This strategy could be used in future fordelivery of DNAzymes into cells [99].

FNA-Based Sensor Against Toxins

Apart from heavy metals, several other toxin molecules (inorganic, organic, and biological)exist in our daily environment. In firmasector, there are oodles of misuses of pesticides,insecticides (organochlorines, organophosphates, carbamates, organochlorine like DDT, etc.)which are mostly hazardous to its surrounding niche. The farmers, firming animals, and eventhe common people suffer frequently due to these toxic contaminants. In addition, there are somany biological agents (microbes, pathogens, toxins) which are tiny to detect but enormous inits adverse effect. Various traditional detection methods are available which have limitations inits sensitivity and specificity. To address all these issues, modern trends in sensor design relyon functional nucleic acids as a smart sensing agent. A few aptamer-based sensors fordetection and monitoring of environmental contaminants had already been discussed in ourprevious review [100].

In a study, scientists had developed a simple aptasensor for sensitive and selective detectionof acetamiprid which is an odorless neonicotinoid insecticide. This has been developed onbasis of electrochemical impedance spectroscopy (EIS). To enhance sensitivity of theaptasensors, AuNPs were electrodeposited on Au electrode surface by cycle voltammetry(CV), which was employed as a platform for aptamer immobilization. With the addition ofacetamiprid, the formation of acetamiprid–aptamer complex on the AuNPs-deposited electrodesurface resulted in an increase of electron transfer resistance. The detection limit is as low as1 nM of acetampirid [101]

Recently, a chemiluminescence competitive aptamer assay has been developed againstaflatoxin B1 (AFB1), a mycotoxin using a hemin/G-quadruplex horseradish peroxidase-mimicking DNAzyme (HRP-DNAzyme) linked with AFB1-specific aptamer candidate[102]. This novel approach exhibited a wide dynamic range from 0.1 to 10 ng/mL with alimit of detection of 0.11 ng/mL and showed no cross reactivity with other mycotoxinsorherbicides like atrazine.

Ochratoxin A (OTA) is another mycotoxin which commonly contaminate foods, particu-larly cereals and cereal products, with strict low regulatory levels (of ppb) in many countriesworldwide. An electrochemical competitive aptamer-based biosensor for OTA is described.


Paramagnetic microparticle beads (MBs) were functionalized with an aptamer specific toOTA, and the magnetic aptasensors showed a linear response to OTA in the range 0.78–8.74 ng/mL and a limit of detection of 0.07±0.01 ng/mL [103].

Polychlorinated biphenyls (PCBs) are one of major concerns for decades due to theirpotential toxicity to human health. Many traditional detection methods have been developed.To trace PCBs efficiently and sensitively, aptamers, a new class of diagnostic tools, areconsidered to be such additional candidates for detection of pollutants.

In a study, scientists had invented the novel method for detection of biological agents(spores, airborne microbes) using novel compositions, methods, and apparatus comprising oneor more nucleic acid ligands functionally coupled to an organic semiconductor. More partic-ularly, the invention relates to the production and use of nucleic acid ligands against anthraxspores [104].

Aptamer-based biosensors, which can easily adopt their surroundings, tempt scientists indesigning highly sensitive, selective, and structure switchable sensing assays. Through intri-cate design, chemical functionalization, numerous aptamer-based assays have been developedthat can switch their conformation upon incubation with target analytes, resulting in anenhanced output signal. To further lower the detection limits to picomolar levels, nanomaterialshave attracted great interest in the design of aptamer-based sensing platforms. Associated totheir unique properties, nanomaterials offer great promise for aptasensors-based applications.

Advanced FNA Strategy

The recent trends of FNA sensor technology depend on allosteric DNAzymes or aptazymes.DNAzymes which can perform chemical modifications on nucleic acids, which can be coupledto aptamers “the magical bullet,” can bind to a broad range of molecules. A combination of thetwo has generated a new class of functional nucleic acids known as aptazymes. This hybridstrategy of allosteric ribozyme combines aspect of both the effector-binding and activity-basedselection approaches. Scientists had devised various methods to fabricate novel types ofaptazymes. Aptazymes can be fabricated by integration of aptamers with hammerhead ribo-zyme, hairpin ribozyme, or ligase ribozyme. In several studies, enzymes contain a replaceablestem-loop or hairpin structure where aptamers can be incorporated easily [105]. Engineered“aptazymes” wherein aptamers fused with hammerhead ribozymes are intelligent biosensingcomponent through ligand-induced conformational rearrangement and activation [107].Hammerhead ribozymes are also an integrative part of designing novel aptazymes. In thiscase, the substrate and enzyme binding arm act in transmanner. In addition to the hammerheadribozyme, many other nucleic acid enzymes have also been employed as scaffolds foraptazymes. Ligase ribozyme is another such molecule which can be readily engineered tofunction as an allosteric enzyme and reveal that many of the techniques and principlesdemonstrated during the development of hammerhead aptazymes may be generalizable. Aselected ribozyme ligase, L1, has been engineered to respond to small organic effectors.Aptamers which can bind small molecules like adenosine and theophylline were attached tothe core structure, and the resultant aptazymes are responsive to their cognate effectors.Rational sequence substitutions in the joining region between the aptamer and the ribozymeyielded aptazymes whose activities were enhanced from 800–1,600-fold in the occurrence of1 mMATP or theophylline [105]. There are different strategies available to modulate ribozymestructure along with its substrate binding arm and the location where aptamer ligand will bind(Fig. 6). Since aptazymes are essentially nucleic acid enzymes, the signaling methods availablefor DNAzymes, ribozymes are applicable to aptazyme-based sensors also. Several fluorescent


signaling methods can be incorporated in aptazyme based sensors [106]. Not surprisingly,aptazymes-based sensors are also developed based on colorimetric signaling methods. Theaptazyme was built on the Pb2 + -specific DNAzyme catalytic core, and the substrate wasextended to bind AuNPs [108]. Using L1-ligase-based aptazyme, other platforms like mass-sensitive quartz-crystal microbalance (QCM) was also exploited by Ellington and coworkers[109]. But current trends of aptazyme based sensing machinery lead to genosensor in whichthe target is primarily genetic material. But, recently aptazymes are a promising candidate as ancustom-made, allosteric molecule for sensing environmental pollutants.

Conclusion

Since the invention of functional nucleic acids, they are incessantly exploited as a sensing tool.However, the sensing and diagnostic market is still primarily dominated by antibodies andother diagnostic tools. Numerous nucleic-acid-based detection methods have been introducedto address this deficiency; however, the costs and requirement for expensive equipment maylimit the widespread use of such technologies. Thus, there is an unmet demand of newplatform technology to improve the detection and identification of environment pollutants.Although various FNA based sensors are reported enormously, the in-house strategy suffersfrom on-site validation and other issues. But on the onset of “nanotechnology” era,nanomaterials like carbon nanotubes (CNTs), QDs, graphenes, metal nanoparticles, and metaloxide nanostructures will aid in the development of FNA-based sensors for environmental

Fig. 6 a–c Examples of NAEs with replaceable hairpins (in blue) in the enzyme strand. d–h Examples ofaptazymes based on appending aptamers to stem II of the hammerhead ribozyme. i An aptazyme activated byboth theophyllilne and FMN with cooperativity. Optimization of aptazymes with in vitro selections by random-izing the communication module (j) or the aptamer (k). lAHg2 + -activated aptazyme based on a UO2

2+-specificDNAzyme (Reprinted with permission Liu [106], Copyright © 2009, American Chemical Society)


monitoring. The amalgamation of nanotechnology, material science, and bioanalytical systemswill inevitably lead to a golden future of sensor research. Moreover, in the future, these FNAscan be integrated in more complex and multifaceted integral system micro total analysissystem (μTAS) or lab-on-a-chip (LOC) to detect pollutants on-site. Given the growing demandfor convenient, economical, portable, yet effective sensors, functional nucleic acids will be akey component in sensor world.

References

1. Ligler, F. S. and Taitt, C.R. (2002) A Optical Biosensors: Present and Future Elsevier Science B.V.:Amsterdam.

2. Sadik, O. A., Wanekaya, A. K., & Andreescu, S. (2004). Journal of Environmental Monitoring, 6, 513–522.

3. Tondeur, Y., Niederhut, W. N., Campana, J. E., & Missler, S. R. (1987). Biomedical and EnvironmentalMass Spectrometry, 14(8), 449–456.

4. Breaker, R. R. (1997). Chemistry Review, 97, 371.5. Kopylov, A. M., & Spiridonova, V. A. (2000). Molecular Biology, 34, 940.6. Guerrier-Takada, C., Gardiner, K., Marsh, T., Pace, N., & Altman, S. (1983). Cell, 35, 849–857.7. Symons, R. H. (1992). Annual Review of Biochemistry, 61, 641–671.8. Cech, T. R. (1983). Cell, 34, 713.9. Ban, N., Nissen, P., Hansen, J., Moore, P. B., & Steitz, T. A. (2000). Science, 289, 902.

10. Pan, T., & Uhlenbeck, O. C. (1992). Nature, 358, 560–563.11. Bartel, D. P., & Szostak, J. W. (1993). Science, 261, 1411–1418.12. Ekland, E. H., Szostak, J. W., & Bartel, D. P. (1995). Science, 269, 364–370.13. Chapman, K. B., & Szostak, J. W. (1995). Chemical Biology, 2, 325–333.14. Ekland, E. H., & Bartel, D. P. (1996). Nature, 382, 373.15. Tsukiji, S., Pattnaik, S. B., & Suga, H. (2003). Nature Structural Biology, 10, 713.16. Conn, M. M., Prudent, J. R., & Schultz, P. G. (1996). Journal of the American Chemical Society, 118, 7012.17. Chapple, K. E., Bartel, D. P., & Unrau, P. J. (2003). RNA, 9, 1208.18. Feng, L. Y., Chen, Y., Ren, J. S., & Qu, X. G. (2011). Biomaterials, 32, 2930.19. Kim, Y. J., Kim, Y. S., Niazi, J. H., & Gu, M. B. (2010). Bioprocess Biosyst. Engineering, 33,

31–37.20. Nolte, A., Klussmann, S., Bald, R., Erdmann, V. A., & Fürste, J. P. (1996). Nature Biotechnology, 14(9),

1116–1119.21. Klussmann, S., Nolte, A., Bald, R., Erdmann, V. A., & Fürste, J. P. (1996). Nature Biotechnology, 14(9),

1112–1115.22. Williams, K. P., Liu, X. H., Schumacher, T. N., Lin, H. Y., Ausiello, D. A., Kim, P. S., & Bartel, D. P.

(1997). Proceedings of the National Academy of Sciences, 94(21), 11285–11290.23. Breaker, R. R. (1996). Current Opinion in Biotechnology, 7, 442–448.24. Corey, M. J., & Corey, E. (1996). Proceedings of the National Academy of Sciences, 93, 11428–11434.25. Breaker, R. R., & Joyce, G. F. (1995). Chem. Biol., 2, 655.26. Bruesehoff, P. J., Li, J., Augustine, A. J., & Lu, Y. (2002). Comb. Chem., 5, 327.27. Joyce, G. F. (2004). Annu. Rev. Biochem., 73, 791–836.28. Silverman, S. K. (2005). Nuclear Acids Research, 33(19), 6151–6163.29. Robertson, M. P., Hesselberth, J. R., & Ellington, A. D. (2001). RNA, 7(4), 513–523.30. Purschke, W. G., Radtke, F., Kleinjung, F., & Klussmann, S. (2003). Nuclear Acids Research, 31(12),

3027–3032.31. Duffus, J. H. (2002). “Heavy metals” a meaningless term? (IUPAC Technical Report). Pure and Applied

Chemistry, 74, 793–807.32. Klopman, G. (1974). Generalized perturbation theory of chemical reactivity, Chemical Reactivity and

Reaction Paths, New York (pp. 55–165). USA: John Wiley.33. Frausto da Silva, J. J. R., & Williams, R. J. P. (1993). The biological chemistry of the elements: the

inorganic chemistry of life. Oxford, UK: Oxford University Press.34. Nieboer, E., & Richardson, D. H. S. (1980). Environmental Pollution (Series B), 1, 3–26.35. Gloag, D. (1981). British Medical Journal (Clinical Research Ed.), 282, 41–44.36. Liu, J., & Yi, L. (2003). Journal of the American Chemical Society, 125, 6642–6643.37. Liu, J., & Yi, L. (2004). Chemistry of Materials, 16, 3231–3238.


38. Liu, J., & Yi, L. (2004). Journal of the American Chemical Society, 126, 12298–12305.39. Swearingen, C. B., Wernette, D. P., Cropek, D. M., Lu, Y., Sweedler, J. V., & Bohn, P. W. (2005). Analytical

Chemistry, 77, 442–448.40. Chang, I. H., Tulock, J. J., Liu, J., Kim, W. S., Cannon, D. M., Jr., Lu, Y., Bohn, P. W., Sweedler, J. V., &

Cropek, D. M. (2005). Environmental Science and Technology, 39, 3756–3761.41. Wernette, D. P., Swearingen, C. B., Cropek, D. M., Lu, Y., Sweedler, J. V., & Bohn, P. W. (2006). Analyst,

131, 41–7.42. Dalavoy, T. S., Wernette, D. P., Gong, M., Sweedler, J. V., Lu, Y., Flachsbart, B. R., Shannon, M. A., Bohn,

P. W., & Cropek, D. M. (2008). Lab Chip, 8, 786–93.43. Chen, X., Guan, H. L., He, Z. K., Zhou, X. D., & Hu, J. M. (2012). Analysis Methods, 4, 1619–1622.44. Shen, L., Chen, Z., Li, Y., He, S., Xie, S., Xu, X., Liang, Z., Meng, X., Li, Q., Zhu, Z., Li, M., Le, X. C., &

Shao, Y. (2008). Analytical Chemistry, 4, 6323–6328.45. Mazumdar, D., Liu, J., Lu, G., Zhou, J., & Lu, Y. (2010). Chemical Communications, 46, 1416–1418.46. Wang, F., Wu, Z., Lu, Y., Wang, J., Jiang, J. H., & Yu, R. Q. (2010). Annual Biochemistry, 405, 168–73.47. Guo, L., Nie, D., Qiu, C., Zheng, Q., Wu, H., Ye, P., Hao, Y., Fu, F., & Chen, G. (2012). Biosensors and

Bioelectronics, 35, 123–7.48. Zhao, Y., Zhang, Q., Wang, W., & Jin, Y. (2013). Biosensors and Bioelectronics, 43, 231–236.49. Gao, A., Tang, C. X., He, X. W., & Yin, X. B. (2013). Analyst, 138, 263–268.50. Li, T., Wang, E., & Dong, S. (2010). Analytical Chemistry, 82, 1515–1520.51. Pelossof, G., Tel-Vered, R., & Willner, I. (2012). Analytical Chemistry, 84, 3703–3709.52. Zhuang, J., Fu, L., Xu, M., Zhou, Q., Chen, G., & Tang, D. (2013). Biosensors and Bioelectronics, 45, 52–

7.53. Nie, D., Wu, H., Zheng, Q., Guo, L., Ye, P., Hao, Y., Li, Y., Fu, F., & Guo, Y. (2012). Chemical

Communications, 48, 1150–1152.54. Li, H., Zhang, Q., Cai, Y., Kong, D. M., & Shen, H. X. (2012). Biosensors and Bioelectronics, 34, 159–

164.55. Smirnov, I., & Shafer, R. H. (2000). Journal of Molecular Biology, 296, 1–5.56. Brenneman, K. L., Poduri, S., Stroscio, M. A., & Dutta, M. (2013). IEEE Sensors Journal, 13, 1783–1786.57. Nicol, C. W. (2012). Minamata: a saga of suffering and hope”Japan times, 7, 10.58. Ministry of the Environment, Govt of Japan, Minamata Disease The History and Measures; Chapter 259. Bakir, F., Rustam, H., Tikriti, S., Al-Damluji, S. F., & Shihristani, H. (1980). Postgraduate Medical

Journal, 56, 1–10.60. Carvalho, C. M., Chew, E. H., Hashemy, S. I., Lu, J., & Holmgren, A. (2008). Journal of Biological

Chemistry, 283, 11913–23.61. ATSDR - Mercury - Regulations and Advisories, p509-524.62. Braman, R. (1971). Analytical Chemistry, 43, 1462–1467.63. Ono, A., & Togashi, H. (2004). Angewandte Chemie International Edition in English, 43, 4300–4302.64. Liu, J., & Lu, Y. (2007). Angewandte Chemie International Edition in English, 46, 7587–7590.65. Chiang, C. K., Huang, C. C., Liu, C. W., & Chang, H. T. (2008). Analytical Chemistry, 80, 3716–3721.66. Wang, J., & Liu, B. (2008). Chemical communications (Cambridge), 39, 4759–4761.67. Liu, B. (2008). Biosensors and Bioelectronics, 24, 762–766.68. Xue, X., Wang, F., & Liu, X. (2008). Journal of the American Chemical Society, 130, 3244–3245.69. Li, D., Wieckowska, A., & Willner, I. (2008). Angewandte Chemie International Edition in English, 47,

3927–3931.70. Wu, D., Zhang, Q., Chu, X., Wang, H., Shen, G., & Yu, R. (2010). Biosensors and Bioelectronics, 25,

1025–1031.71. Xu, X., Wang, J., Jiao, K., & Yang, X. (2009). Biosensors and Bioelectronics, 24, 3153–3158.72. Yu, C. J., Cheng, T. L., & Tseng, W. L. (2009). Biosensors and Bioelectronics, 25, 204–210.73. Li, T., Dong, S., & Wang, E. (2009). Analytical Chemistry, 81, 2144–2149.74. Li, T., Li, B., Wang, E., & Dong, S. (2009). Chemistry of Communications (Cambridge), 24, 3551–3553.75. Hoang, C. V., Oyama, M., Saito, O., Aono, M., & Nagao, T. (2013). Science Reports, 3, 1175.76. Brewer, G. J. (2010). Clinical Neurophysiology, 121, 459–460.77. Casarett & Doull’s Toxicology, The basic science of poisons, Fifth Edition, Edited by Curtis D. Klassen,

McGraw-Hill, New York. pp 715.78. Edelstein DL. U.S. Geological Survey, Mineral commodity summaries, January 2013.79. Environmental Protection Agency, National Primary Drinking Water Regulations for Lead and Copper,

Federal Register / Vol. 65, No. 8 / Wednesday, January 12, 2000 / Rules and Regulations. pp. 1950-2015.80. Carmi, N., Shultz, L. A., & Breaker, R. R. (1996). Chemical Biology, 3, 1039–1046.81. Carmi, N., Balkhi, S. R., & Breaker, R. R. (1998). Proceedings of the National Academy of Sciences, 95,

2233–2237.


82. Carmi, N., & Breaker, R. R. (2001). Bioorganic and Medicinal Chemistry, 9, 2589–2600.83. Liu, J., & Lu, Y. (2007). Journal of the American Chemical Society, 129, 9838–9839.84. Zuo, P., Yin, B. C., & Ye, B. C. (2009). Biosensors and Bioelectronics, 25, 935–939.85. Wang, Y., Yang, F., & Yang, X. (2010). Nanotechnology, 21, 205502.86. Su, Y. T., Lan, G. Y., Chen, W. Y., & Chang, H. T. (2010). Analytical Chemistry, 82, 8566–8572.87. Miao, X., Ling, L., Cheng, D., & Shuai, X. (2012). Analyst, 137, 3064–3069.88. Fang, Z., Huang, J., Lie, P., Xiao, Z., Ouyang, C., Wu, Q., Wu, Y., Liu, G., & Zeng, L. (2010). Chemical

communications (Cambridge), 46, 9043–9045.89. Zhang, L., Zhu, J., Ai, J., Zhou, Z., Jia, X., & Wang, E. (2013). Biosensors and Bioelectronics, 39, 268–

273.90. Yin, B. C., Ye, B. C., Tan, W., Wang, H., & Xie, C. C. (2009). Journal of the American Chemical Society,

131, 14624–14625.91. Chen, Z., Li, L., Mu, X., Zhao, H., & Guo, L. (2011). Talanta, 85, 730–735.92. Lin, Z., Li, X., & Kraatz, H. B. (2011). Analytical Chemistry, 83, 6896–6901.93. Shi, L., Liang, G., Li, X. H., & Liu, X. H. (2012). Analytical Methods, 4, 1036–1040.94. Adriaens P, Vannela R, (2010) Dnazymes and sensors incorporating the Same, US Patent No - US 7,709,

619 B2.95. Wu, Y., Zhan, S., Wang, F., He, L., Zhi, W., & Zhou, P. (2012). Chemical communications (Cambridge),

48, 4459–4461.96. Wu, Y., Liu, L., Zhan, S., Wang, F., & Zhou, P. (2012). Analyst, 137, 4171–4178.97. Liu, J., Brown, A. K., Meng, X., Cropek, D. M., Istok, J. D., Watson, D. B., & Lu, Y. (2007). Proceedings

of the National Academy of Science, 104, 2056–2061.98. Tang, Q., Yuan, Y., Xiao, X., Guo, P., Hu, J., Ma, D., & Gao, Y. (2013). Microchimica Acta, 180, 1059–

1064.99. Wu, P., Hwang, K., Lan, T., & Lu, Y. (2013). Journal of the American Chemical Society, 135, 5254–5257.100. Sett, A., Das, S., Sharma, P., & Bora, U. (2012). Open Journal of Applied Biosensors, 1, 9–19.101. Fana, L., Zhaoa, G., Shia, H., Liua, M., & Lia, Z. (2013). Biosensors and Bioelectronics, 43, 12–18.102. Shim, W. B., Mun, H., Joung, H.-A., Ofori, J. A., Chung, D.-H., & Kim, M. G. (2014). Food Control,

36(1), 30–35.103. Bonel, L., Vidal, J. C., Duato, P., & Castillo, J. R. (2011). Biosensors and Bioelectronics, 26(7), 325–9.104. Vivekananda, J.(2003). Methods and compositions for aptamers against anthrax. US Patent no- US6569630

B1.105. Robertson, M. P., & Ellington, A. D. (2000). Nuclear Acids Research, 28(8), 1751–1759.106. Liu, J., Cao, Z., & Lu, Y. (2009). Chemistry Reviews, 109, 1948–1998.107. Silva, C., & Walter, N. G. (2009). RNA, 15(1), 76–84.108. Liu, J., & Lu, Y. (2004). Analytical Chemistry, 76, 1627–32.109. Knudsen, S. M., Lee, J., Ellington, A. D., & Savran, C. A. (2006). Journal of the American Chemical

Society, 128(50), 15936–15937.


Documents

Functional Nucleic-Acid-Based Sensors for Environmental Monitoring