DEVELOPMENT OF A NANOPARTICLE-BASED ELECTROCHEMICAL BIO-BARCODE
DNA BIOSENSOR FOR MULTIPLEXED PATHOGEN DETECTION ON SCREENPRINTED CARBON ELECTRODES
By
Deng Zhang

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Biosystems Engineering
2011

ABSTRACT
DEVELOPMENT OF A NANOPARTICLE-BASED ELECTROCHEMICAL BIO-BARCODE
DNA BIOSENSOR FOR MULTIPLEXED PATHOGEN DETECTION ON SCREENPRINTED CARBON ELECTRODES
By
Deng Zhang
A highly amplified, nanoparticle-based, bio-barcoded electrochemical biosensor for the
simultaneous multiplexed detection of the protective antigen A (pagA) gene (accession number =
M22589) from Bacillus anthracis and the insertion element (Iel) gene (accession number =
Z83734) from Salmonella Enteritidis was developed. The biosensor system is mainly composed
of three nanoparticles: gold nanoparticles (AuNPs), magnetic nanoparticles (MNPs), and
nanoparticle tracers (NTs), such as lead sulfide (PbS) and cadmium sulfide (CdS). The AuNPs
are coated with the first target-specific DNA probe (1pDNA), which can recognize one end of
the target DNA sequence (tDNA), and many NT-terminated bio-barcode ssDNA (bDNA-NT),
which act as signal reporter and amplifier. The MNPs are coated with the second target-specific
DNA probe (2pDNA) that can recognize the other end of the target gene. After binding the
nanoparticles with the target DNA, the following sandwich structure is formed: MNP2pDNA/tDNA/1pDNA-AuNP-bDNA-NTs. A magnetic field is applied to separate the sandwich
structure from the unreacted materials. Because the AuNPs have a large number of nanoparticle
tracers per DNA probe binding event, there is substantial amplification. After the nanoparticle
2+

tracer is dissolved in 1 mol/L nitric acid, the NT ions, such as Pb

2+

and Cd , show distinct non-

overlapping stripping curves by square wave anodic stripping voltammetry (SWASV) on screenprinted carbon electrode (SPCE) chips. The oxidation potential of NT ions is unique for each
nanoparticle tracer and the peak current is related to the target DNA concentration. The results

show that the biosensor has good specificity, and the sensitivity of single detection of pagA gene
from Bacillus anthracis using PbS NTs is as low as 0.2 pg/mL. The detection limit of this
multiplex bio-barcoded DNA sensor is 50 pg/mL using PbS or CdS NTs. The nanoparticle-based
bio-barcoded DNA sensor has potential applications for multiple detections of bioterrorism
threat agents, co-infection, and contaminants in the same sample.

© Copyright By
Deng Zhang
2011

ACKNOWLEDGMENTS

First, I would like to thank my wife, Yaling Liu, for her unconditional love and support
during the most important period of my life. Her love and trust provide me the power to
overcome all difficulties in the research and life.
I would like to express my sincere appreciation to Dr. Evangelyn Alocilja, for her constant
support, guidance, encouragement, patience, and funding for the last five years. It is my great
honor to have her as my mentor and friend. I am also grateful for the advice and resource
provided by Dr. John Gerlach, Dr. Daniel Guyer, and Dr. Gary Blanchard.
I would like to thank my colleagues in the lab, Zarini Muhammad-Tahir, Sudeshna Pal,
Yang Liu, Edith Torres-Chavolla, Yilun Luo, Michelle Packard, Shannon K. McGraw, Michael
Anderson, and Yun Wang, for their help and motivation. I also want to thank those
undergraduates and high school students who worked with me over the past years.

v

TABLE OF CONTENTS

LIST OF TABLES ---------------------------------------------------------------------------------------------- viii
LIST OF FIGURES ------------------------------------------------------------------------------------ ix
CHAPTER 1. INTRODUCTION
1.1 Objective and Goals --------------------------------------------------------------------------------------1
1.2 Foodborne Disease and Bioterrorism --------------------------------------------------------------------- 2
1.3 Current Detection Methods--------------------------------------------------------------------------------- 4
1.4 Use of Biosensors-------------------------------------------------------------------------------------------- 6
1.5 Novelty of Research----------------------------------------------------------------------------------------11
1.6 Hypothesis ---------------------------------------------------------------------------------------------------12
1.7 Specific Aims------------------------------------------------------------------------------------------------12
CHAPTER 2. LITERATURE REVIEW
2.1 Overview of Nano-sensors ------------------------------------------------------------------ 15
2.2 Electrochemical DNA Sensor --------------------------------------------------------------- 24
2.3 Bio-barcode Assay --------------------------------------------------------------------------------- 29
2.4 Screen-printed Carbon Electrode ---------------------------------------------------------------- 31
2.5 Square Wave Anodic Stripping Voltammetry-------------------------------------------------- 32
CHAPTER 3. MATERIALS AND METHODS
3.1 Reagents and Materials ----------------------------------------------------------------------- 36
3.2 Facilities and Equipment --------------------------------------------------------------------- 37
3.3 Synthesis of Nanoparticles ------------------------------------------------------------------- 38
3.4 Bacteria Culture, DNA isolation, PCR and DNA Probes --------------------------------- 39
3.5 Functionalization of Nanoparticles ---------------------------------------------------------- 40
3.6 Confirmation of Nanoparticle Functionalization ------------------------------------------ 44
3.7 Validation and Optimization of the Biosensor --------------------------------------------- 47
3.8 Electrochemical Analysis--------------------------------------------------------------------- 47
CHAPTER 4. RESULTS AND DISCUSSION
4.1 Characterization of Nanoparticles ----------------------------------------------------------- 49
4.2 Functionalization of Nanoparticles ---------------------------------------------------------- 51
4.3 Fluorescent Detection Using Bio-barcode Assay ------------------------------------------ 54
4.4 Electrochemical Validation and Characterization of SPCE Chips----------------------- 56
4.5 Detection of AuNPs on SPCE Chips -------------------------------------------------------- 63
4.6 Biosensor Detection of Single Target DNA ------------------------------------------------ 67
4.7 Biosensor Detection of Multiple Target DNA -------------------------------------------- 70
4.8 Conclusion and Summary------------------------------------------------------------------------- 75
APPENDICES----------------------------------------------------------------------------------------------- 78
Appendix I. DNA Extraction Procedure -------------------------------------------------------- 79
Appendix II. PCR Amplification ---------------------------------------------------------------- 81
vi

Appendix III. Procedure of Bio-Barcode Assay------------------------------------------------ 82
Appendix IV. Buffer Preparation ---------------------------------------------------------------- 87
Appendix V. Conjugation of carboxylic group on NTs with amine group on bio-barcoded
AuNPs --------------------------------------------------------------------------------------------------- 88
BIBLIOGRAPHY ------------------------------------------------------------------------------------------ 89

vii

LIST OF TABLES

Table 1.1 Foodborne diseases in the US by major foodborne pathogens in 2006 (CDC 2010).--- 2
Table 2.1. Electrochemical genosensors using AuNPs as label. GE: gold electrode; DPV:
differential pulse voltammetry; ASV: anodic stripping voltammetry; SPMBE: sandwich-type
screen-printed microband electrode; PSA: potentiometric stripping analysis; SPEs: screenprinted electrodes; M-GECE: magnetic graphite-epoxy composite electrode, GCE: glassy carbon
electrode (Castada, Alegret et al. 2007). ----------------------------------------------------------------- 19
Table 2.2 Comparison of platforms for DNA electrochemical sensing (Drummond, Hill et al.
2003). --------------------------------------------------------------------------------------------------------- 26
Table 4.1. Fluorescence signal of supernatant solution for evaluation of conjugation efficiency
between MNPs and 2pDNA. ------------------------------------------------------------------------------ 51
Table 4.2. The relationship between target DNA and the fluorescence signal of released bDNA.55
Table 5. Buffer preparation. ------------------------------------------------------------------------------- 87

viii

LIST OF FIGURES

Figure 1.1. Schematic of a biosensor. (A) Biological reaction between biological sensing element
and analytes; (B) Transduction from biological event to electrical output. For interpretation of
the references to color in this and all other figures, the reader is referred to the electronic version
of this dissertation.------------------------------------------------------------------------------------------- 7
Figure 1.2. Schematic of a three-electrode electrochemical detection system. The system is
composed of a potentiostat, working, reference, and auxiliary electrodes. (adapted from Bard
and Faulkner 2001). ----------------------------------------------------------------------------------------- 9
Figure 1.3. Schematic of DNA base pairs.--------------------------------------------------------------- 10
Figure 2.1. The different strategies used for the integration of gold nanoparticles (AuNPs) into
DNA-sensing systems: (A) previously dissolving AuNPs by using HBr/Br2 mixture followed by
Au(III)-ion detection; (B) direct detection of AuNPs anchored onto the surface of the genosensor;
(C) conductometric detection; (D) enhancement with Ag or Au followed by detection; (E)
AuNPs as carriers of other AuNPs; and, (F) AuNPs as carriers of other electroactive labels
(adapted from Castada, Alegret et al. 2007). ------------------------------------------------------------ 17
Figure 2.2. Magnetic beads (MBs) as versatile tools for bioassays. Beads modified with various
recognition elements (examples of commercially available MBs shown on the gray background)
can be used for specific bioaffinity capture of different molecules. For example, beads bearing
covalently attached oligo(dT)25 chains can bind nucleic acid involving (A)n stretches, including
natural eukaryotic mRNAs or tDNAs tagged with (A)n adaptors. Streptavidin-coated beads are
suitable for capturing any biotinylated molecules, including single strand or double strand
nucleic acids, aptamers, peptides, proteins, etc. Antibodies can be attached to the beads either
via a direct covalent linkage, or via specific antibody binding proteins such as protein A or
protein G (beads functionalized with the latter proteins can be further modified with various
antibodies on demand). Other affinity ligands attached to MBs are suitable for specific capture
of tagged recombinant proteins (such as metal-affinity cobalt chelate for histidine tags).
Biomolecules captured at the MB surface can serve as a biorecognition layer for interaction with
other molecules. For example, a biotinylated single strand oligonucleotide (ssODN) immobilized
at streptavidin-coated beads features a hybridization probe for complementary sequence, the
dsDNA can represent a target for a DNA-binding protein, antibody can bind a specific antigen
(including antigens exposed at surfaces of whole cells), immobilized protein (e.g., via antibody,
his-tagor biotin–streptavidin linkage) can interact with a specific nucleic acid or another protein,
etc. (adapted from Palecek and Fojta 2007). ------------------------------------------------------------ 20
Figure 2.3. Schematic of an electrochemical DNA biosensor. (A) DNA probe immobilization; (B)
hybridization between DNA probe and target DNA on the transducer; (C) signal processing by a
biosensor system. ------------------------------------------------------------------------------------------- 25

ix

Figure 2.4. Multi-target electrical DNA detection protocol based on different inorganic colloid
nanoparticle tracers. (A) Introduction of probe-modified magnetic beads. (B) Hybridization with
the DNA targets. (C) Second hybridization with the QD-labeled probes. (D) Dissolution of QDs
and electrochemical detection (adapted from Wang, Liu et al. 2003). ------------------------------- 28
Figure 2.5. Schematic of bio-barcode DNA assay. (A) formation of MNP-2pDNA
/tDNA/1pDNA-AuNP-bDNA sandwich structure; (B) probe separation and bDNA release. ---- 30
Figure 2.6. Waveform and measurement scheme for square wave voltammetry. Shown in bold is
the actual potential waveform applied to the working electrode. The light intervening lines
indicate the underlying staircase onto which the square wave can be regarded as having been
superimposed. In each cycle, a forward current sample is taken at the time indicated by the solid
dot, and a reverse current sample is taken at the time marked by the shaded dot (Adapted from
Bard and Faulkner 2000). ---------------------------------------------------------------------------------- 33
Figure 2.7. Principle of anodic stripping. Values shown are typical ones used; potentials and Ep
2+
are typical of Cu analysis, (a) Preelectrolysis at Ed; stirred solution, (b) Rest period; stirrer off.
(c) Anodic scan (v = 10-100 mV/s) (Adapted from Bard and Faulkner 2000). -------------------- 34
Figure 3.1. Schematic of screen-printed carbon electrode: A) top view, B) cross-section view. It
is composed of working electrode (carbon electrode) and auxiliary/reference electrode (Ag/AgCl)
in an insulating well.---------------------------------------------------------------------------------------- 38
Figure 3.2. Schematic of the functionalization of AuNPs. A mixture of thiolated
oligonucleotides (bDNA:1pDNA=100:1) form a self assembled monolayer on the surface of
gold nanoparticles. ---------------------------------------------------------------------------------------- 41
Figure 3.3. Schematic of the functionalization of MNPs. (1) formation of maleimide-activated
MNPs; (2) formation of MNP-2pDNA conjugate. ----------------------------------------------------- 42
Figure 3.4 Evaluation of conjugation efficiency: MNP- pDNA and AuNP-bDNA. --------------- 43
Figure 3.5. Schematic of conjugation of bio-barcoded AuNP with NTs. (1) functionalization of
gold nanoparticles with thiolated bDNA and thiolated 1pDNA; (2) formation of the 1pDNAAuNP-bDNA-NTs complex by cross-linking amine group on bDNA and carboxylic group on
NTs through 1-Ethyl-3-[3-dimethylaminopropyl] carbodiimide Hydrochloride (EDC) and Nhydroxysuccinimide (NHS).------------------------------------------------------------------------------- 44
Figure 3.6. Schematic of the bio-barcode assay: A) formation of MNP-2 pDNA/target
DNA/1pDNA-AuNP-bDNA-fluorescein; B) bDNA-fluorescein separation and release.--------- 45
Figure 3.7. Schematics of electrochemical measurement of AuNPs. It is composed of 2 steps:
3+
AuNP dissolution on the SPCE and DPV measurement of Au .------------------------------------ 46
Figure 3.8. The composition of the electrochemical detection system. ------------------------------ 48

x

Figure 4.1. Characterization of AuNPs: A) Transmission electron microscopy (TEM) image of
AuNPs, showing an average of 15 nm in diameter; B) Absorbance spectrum of AuNPs at 519 nm.
----------------------------------------------------------------------------------------------------------------- 49
Figure 4.2. TEM images of NTs: A) CdS, showing an average diameter of 7 nm; B) PbS,
showing an average diameter of 3 nm. ------------------------------------------------------------------- 50
Figure 4.3. Fluorescence signal of supernatant solution for evaluation of conjugation efficiency
between AuNPs and bDNA. ------------------------------------------------------------------------------ 52
Figure 4.4. Confirmation of the conjugation of NTs and AuNPs. Three test tubes labeled A, B, C
are under white light and UV light after centrifugation at 13,000 rpm for 30 min. (Tube A: biobarcoded AuNPs, NTs, EDC, and NHS; tube B: bio-barcoded AuNPs and NTs; tube C: NTs
only).---------------------------------------------------------------------------------------------------------- 53
Figure 4.5. The fluorescence signal of released bDNA with different concentrations of target
DNA. --------------------------------------------------------------------------------------------------------- 55
Figure 4.6. Cyclic voltammogram of bare SPCE sensor in a 5mM K3Fe(CN)6 solution with 1M
KCl. Scan rate is 50mV/s. --------------------------------------------------------------------------------- 56
Figure 4.7. The repeatability test of 1 mg/L of bismuth film-coated SPCE sensor and bare SPCE
sensor. Deposition: -1.2 V for 10 min; SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz,
amplitude 25 mV, and potential step 5 mV, at room temperature and in nonstirred solution. --- 57
Figure 4.8. Stripping voltammogram of 5 mg/L of lead(II) with various concentrations of
bismuth on electrochemical sensor after 2 min deposition at -1.2 V. SWASV scan from -1.2 V
to 0.0 V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV, at room temperature and
in nonstirred solution. -------------------------------------------------------------------------------------- 59
Figure 4.9. Stripping voltammogram on bismuth film-coated SPCE sensors with various
deposition times. Deposition at-1.2 V; SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz,
amplitude 25 mV, and potential step 5 mV, at room temperature and in nonstirred solution. --- 60
Figure 4.10. Stripping voltammogram of a mixture of lead(II) and cadmium(II) on bismuth filmcoated SPCE sensors (a: 5 mg/L lead(II) and 5 mg/L cadmium(II); b: 2 mg/L lead(II) and 2 mg/L
cadmium(II); c: 0.5 mg/L lead(II) and 0.5 mg/L cadmium(II)). Deposition: -1.2 V for 10 min;
SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV,
with 1mg/mL bismuth, at room temperature and in nonstirred solution. --------------------------- 61
Figure 4.11. Stripping voltammogram of a mixture of lead(II) and cadmium(II) on bare SPCE
sensors. (a: 5 mg/L lead(II) and 5 mg/L cadmium(II); b: 2 mg/L lead(II) and 2 mg/L cadmium(II);
c: 0.5 mg/L lead(II) and 0.5 mg/L cadmium(II)). Deposition: -1.2 V for 10 min; SWASV scan
from -1.2 V to 0.0 V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV, with 1mg/mL
bismuth, at room temperature and in nonstirred solution. --------------------------------------------- 62

xi

Figure 4.12. The peak currents of stripping voltammogram with different concentrations of
lead(II) and cadmium(II). ---------------------------------------------------------------------------------- 63
Figure 4.13. Cyclic voltammogram of unmodified AuNPs (1.17 x 10-8 M) on SPCE : (A) AuNPs
in 0.1M HCl; (B) 0.1 M HCl. CV scan from 1.4V to 0.0, scan rate 100 mV/s after 1.25 V
electrooxidation for 2 min.--------------------------------------------------------------------------------- 64
Figure 4.14. Cyclic voltammogram of probe-modified AuNPs in 0.1 M HCl on SPCE: (A)
accumulation for 20 min; (B) accumulation for 10 min; (C) no accumulation; (D) control, no
AuNPs and no accumulation. Conditions: electrooxidation potential, +1.25 V; electrooxidation
time, 2 min, CV scan from +1.4 V to 0 V, scan rate 100 mV/s, nonstirred solution. -------------- 66
Figure 4.15. DPV hybridization response of different concentrations of DNA targets on SPCE (a:
700 ng/mL; b: 70 ng/mL; c: 7 ng/mL; d: 0.7 ng/mL) and the calibration plot between peak
current and the tDNA concentration (inset). DPV scan from +1.25 V to 0 V, step potential 10
mV, modulation amplitude 50 mV, scan rate 50 mV/s, and nonstirred solution.------------------- 67
Figure 4.16. Schematic of the bio-barcode assay. (A) formation of MNP-2pDNA /tDNA/1
pDNA-AuNP-bDNA/NTs sandwich structure; (B) tDNA separation and tracer dissolution. ---- 68
Figure 4.17. Single detection of various concentrations of pagA gene from Bacillus anthracis
with the bio-barcode biosensor using PbS NTs. (A) stripping voltammogram of
various
concentrations of pagA gene from Bacillus anthracis; (B) the relationship of tDNA concentration
and peak current. Conditions: deposition at -1.2 V for 10 min; SWASV scan from -1.2 V to 0.0
V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV, with 1mg/mL bismuth, at room
temperature and in nonstirred solution. ------------------------------------------------------------------ 69
Figure 4.18. Schematic of the multiple bio-barcode biosensor: A) bio-barcode assay; B) SWASV
measurement. ------------------------------------------------------------------------------------------------ 71
Figure 4.19. Specificity test of the multiple bio-barcode biosensor using distilled water as control.
(A) control and 0.05 μg/mL of the insertion element (Iel) gene of Salmonella Enteritidis; (B)
control and 0.05 μg/mL of pagA gene from Bacillus anthracis; (C) control and a mixture of 0.05
μg/mL of the insertion element (Iel) gene and 0.05 μg/mL of pagA gene. Conditions: deposition
at -1.2 V for 10 min; SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz, amplitude 25 mV,
and potential step 5 mV, with 1mg/mL bismuth, at room temperature and in nonstirred solution.72
2+

2+

Figure 4.20. The stripping signal of released NT ions (Pb
and Cd ) with different
concentrations of tDNAs: A) stripping voltammogram of various concentrations of tDNA; B) the
calibration plot between peak current and the tDNA concentration. --------------------------------- 74
Figure 4.21. Schematic of portable electrochemical biosensor. -------------------------------------- 77

xii

CHAPTER 1: INTRODUCTION

1.1 Objective and Goals
The long term goal of this research is to develop a handheld multiplex DNA sensor for a
rapid, highly specific and sensitive, cost-effective detection of foodborne pathogens, especially
in resource-limited settings or clinical facilities. This is to be accomplished by using a new
biosensor architecture, which combines screen-printed carbon electrode (SPCE) chips, biobarcode assay and a portable potentiostat. The model agents for this proof of concept research
are Salmonella Enteritidis and Bacillus anthracis. The biosensor, designed to detect the
protective antigen A (pagA) gene (accession number = M22589) from Bacillus anthracis and the
insertion element (Iel) gene (accession number = Z83734) from Salmonella Enteritidis, is mainly
composed of three nanoparticles: gold nanoparticles (AuNPs), magnetic nanoparticles (MNPs),
and nanoparticle tracers (NTs, such as PbS and CdS). After hybridization, the nanoparticle tracer
is dissolved in 1 M nitric acid. The NT ions show an attractive non-overlapping stripping
behavior by square wave anodic stripping voltammetry (SWASV) on SPCE chips. Combined
with a battery-powered, handheld potentiostat, the whole biosensor detection system can provide
a rapid, on-site, cost-effective detection for health care professionals, food safety monitoring
personnel, and anti-bioterrorism agents.
The short term goal of this research is to construct a prototype biosensor. The biosensor will
be evaluated for its ability to detect the two target genes specifically and sensitively in the same
liquid media. It represents an innovative approach to detect multiple pathogens with high
sensitivity due to multiple NT ion detection capability with SWASV on SPCE chips, magnetic
extraction and concentration, and inherently amplified signal from the bio-barcode assay.

1

1.2 Foodborne disease and Bioterrorism
Foodborne disease is caused by consuming contaminated foods or beverages. More than 250
different foodborne diseases have been described. Most of these diseases are infections, caused
by a variety of bacteria, viruses, and parasites that can be foodborne. Centers for Disease Control
and Prevention (CDC) reported each year an estimated 76 million cases of foodborne illness
occur in the United States. Over 325,000 hospitalizations and 9,000 deaths are associated with
foodborne diseases each year (CDC 2010).
The five food categories, excluding multi-ingredient foods, linked to the most foodborne
illness outbreaks are seafood, produce, poultry, beef, and eggs. These five food categories were
responsible for 59 % of all outbreaks and 54 % of the illnesses (CDC 2010). Bacterial pathogens
are responsible for 60 % of outbreaks, while viruses cause 24 %, chemicals/toxins cause 15 %,
and parasites cause 1 % (CDC 2010). Although anyone can develop a foodborne illness, the
elderly, young children, the immuno-compromised, pregnant women and their fetuses are most
at risk. Thousands of types of bacteria are naturally present in our environment. The most
prominent bacterial pathogens in the outbreak data are Salmonella, Clostridium, and Escherichia
coli (CDC 2010). Table 1.1 shows the foodborne disease in the US by major foodborne
pathogens in 2006.
Table 1.1 Foodborne diseases in the US by major foodborne pathogens in 2006 (CDC 2010).
Etiology (confirmed or suspected)

Outbreaks Illnesses

Salmonella

117

3296

Clostridium perfringens

34

1880

Escherichia coli, Shiga toxin--producing (STEC)

29

592

Staphylococcus enterotoxin

29

428

2

"Salmonella" bacteria are the most frequently reported cause of foodborne illness. CDC
estimates that 1.4 million people in the United States are infected, and 1,000 people die each year
with salmonellosis. A total of 40666 Salmonella isolates were reported from participating public
health laboratories in 2005 (CDC 2005). The surveillance report from the Food Diseases Active
Surveillance for 2004 identified Salmonella as the most common bacterial infection reported,
accounting for 42 % of the foodborne illness cases in the United States (CDC 2005). There were
117 Salmonella outbreaks in 2006, causing greater than 3,300 illnesses reported to the CDC
Foodborne Outbreak Reporting System. The most common outbreak serotypes is Salmonella
enterica serovar Enteritidis (Salmonella Enteritidis) (CDC 2010). Salmonella Enteritidis is a
major threat for food safety and public health. There is a widespread occurrence of Salmonella
Enteritidis in animals, especially in poultry and swine. Environmental sources of the organism
include water, soil, insects, factory surfaces, kitchen surfaces, animal feces, raw meats, raw
poultry, and raw seafood. People infected with Salmonella Enteritidis develop diarrhea, fever,
and abdominal cramps 12 to 72 hours after infection. In some patients, especially for infants and
young children, pregnant women and their unborn babies, and older adults, Salmonella infection
may spread from the intestines to the blood stream, and then to other body sites and can be lifethreatening unless the person is treated promptly with antibiotics (CDC 2005).
Some foodborne pathogens could potentially be delivered by bioterrorists through foods,
such as Bacillus anthracis. It is a spore forming rod-shaped bacterium and transmissible to
humans through handling or consumption of contaminated animal products. Bacillus anthracis
spores can resist harsh physical and chemical environment, and remain viable in soil for many
years. Infection may also result from inhalation of Bacillus anthracis spores from contaminated
animal products such as wool or the intentional release of spores during a bioterrorist attack.

3

Bacillus anthracis can enter the human body through the intestines (ingestion), lungs (inhalation),
or skin (cutaneous) and causes distinct clinical symptoms based on its site of entry. In 2001,
Bacillus anthracis spores were intentionally distributed through the postal system, causing 22
cases of anthrax, including 5 deaths, and forever changing the realm of public health (Hughes
and Gerberding 2002). Since then, the prospect of bioterrorism in food, water, and agriculture
has identified critical needs in prevention, protection, and mitigation for homeland security.
Rapid, on-site detection or point-of-care diagnosis are one of such needs.
In addition to the health risk associated with contaminated foods, there is the often
devastating economic impact to the food producer. Especially current wide distribution by these
mega-processing plants puts potential outbreak risks on a national and international scale.
Increasing automation in food processing facilities increases the risk of contamination by
6

environmental sources following heat treatment. A recall of 21.7×10 lb of ground beef in 2007
owing to contamination with E. coli O157:H7 resulted in the Topps Meat Company going out of
business after 67 years of operation. Therefore, monitoring of pathogen counts on processing
surfaces is critical in maintaining low or zero counts in food products. The costs of warehousing
along with the potential costs of product recalls have potentially made on-site pathogen testing
economically advantageous.

1.3 Current Detection Methods
The market potential for food pathogen detection was estimated to be $150 million per year.
This study estimated that 144.3 million microbiological tests were conducted in the food industry
in 1999. The majority of these tests came from processed-food plants, followed closely by dairy
plants (Alocilja and Radke 2003).

4

The common detection methods for pathogens include microbiological, immunological and
molecular biological techniques. Microbiological detection often relies on pathogen growth in
culture media, followed by isolation, biochemical identification. For example, Laboratory
procedures for the identification of Bacillus anthracis from CDC include incubation of purified
specimens at 35-37 ℃ for 18-24 h, then the "Medusa head" colony shows the presumptive
identification. For further confirmation, gram stain or india ink stain will be used (Weyant R.S
2001). The microbiological detection of Salmonella in food samples is also laborious and timeconsuming. Several days are required to culture the bacteria from a sample followed by a final
isolation and identification by biochemical and serological techniques. Although microbiological
detection is sensitive and accurate, it is time-consuming and needs special reagents and facilities
in the laboratory.
The immunological detection system, such as enzyme linked immunosorbent assay (ELISA),
utilizes the specific binding reaction between antibody and antigen. For example, a commercial
Salmonella ELISA Test Kit from Bioo Scientific Inc. provides a simple, rapid, sensitive and
cost-effective screening method, which enables government agencies, seafood processors, as
well as quality assurance organizations, to detect positive and negative Salmonella samples to as
5

low as 10 cells/mL level. The ELISA technique is highly specific due to the specificity of
monoclonal antibody. However, its sensitivity is relatively low and hard to satisfy the on-site
detection.
Molecular polymerase chain reaction (PCR) utilizes a combination of reagents and
temperature change schemes to anneal and denature nucleic acid sequences for exponential
amplification of the gene of interest. Quantitative PCR (Q-PCR) is another primer-based
molecular technique that combines the specificity of conventional PCR with the quantitative

5

measurement of fluorescence for determining the presence of specific types of nucleic acid in
environmental samples. Both molecular biological techniques are very sensitive. However, PCR
techniques are often criticized for their complex, expensive, time-consuming, and labor-intensive
procedures and narrow target DNA (tDNA) quantification range after PCR amplification. The
specialized labs staffed by highly trained personnel are also required.
In the meanwhile, some commercial rapid detection methods have been developed, which
use a vast array of tests that include miniaturized biochemical kits, antibody- and DNA-based
tests, and assays that are modifications of conventional tests to speed up analysis. Automation
using traditional detection methods, such as microbiological and immunological methods, is also
increasing in popularity. However, the detection systems are costly, require specialized training,
have complex processing steps, need enrichment of bacteria in an enrichment medium before
analysis. In addition, most of them are only for the detection of one agent. So a rapid, costeffective, sensitive, multiple, user-friendly, on-site detection and valid identification of
pathogenic agents, such as Salmonella Enteritidis and Bacillus anthracis, is vital within the
overall context of food safety, anti-bioterrorism, and point-of-care diagnosis.

1.4 Use of Biosensors
A biosensor is an analytical device that integrates a biological sensing element with a transducer
to quantify a biological event into an electrical output. Figure 1 shows the schematic of a
biosensor. It consists of three components: 1) biological sensing element, which can bind the
biological analytes specifically, such as antibody for antibody, DNA probe (pDNA) for DNA
target (tDNA); 2) transducer, which can convert the biological event into an electrical output,

6

Biological
sensing
element
Analytes

Signal Processing

Transducer

Amplifier
Readout

(A)

(B)

Figure 1.1. Schematic of biosensors. (A) Biological reaction between biological sensing element
and analytes; (B) Transduction from biological event to electrical output. For interpretation of
the references to color in this and all other figures, the reader is referred to the electronic version
of this dissertation.
such as optical fibers and electrodes; 3) associated electronics or signal processors those are
primarily responsible for the display of the results in a user-friendly way. Transduction of
biological reaction is mainly measured electronically (Lee and Shim 2001), optically (Dubertret
2005; Pu, Pan et al. 2008; Sassolas, Leca-Bouvier et al. 2008) or electrochemically (Drummond,
Hill et al. 2003; Weng, Zhang et al. 2008), or using mass-sensitive devices (Huang, Wang et al.
2006).
Optical methods are the most frequently used detection approach currently. The simplest
detection units are spectrometers and fluorometers, which can be used for spectroscopic or
fluorescence detection of target indicators, respectively. Another widely used option is flow
cytometry (FCM), in which cells are physically analyzed based on characteristics such as natural
fluorescence or light scattering. FCM is often paired with immunomagnetic capture to
concentrate cells which are passed single file in a fluid stream with the light scatter from a laser
defining cell count. Advanced flow cytometers can even sort target cells away from waste
materials onto membranes or slides, for further verification methods. FCM systems have been
7

deployed in the field, but they are generally not portable or robust and require advanced training
to operate. Although current optical sensing approaches are effective for high-density arrays,
they are hard to miniaturize and use for in-field detection or point-of-care diagnosis.
Electrochemical systems for biomedical diagnostics can be miniaturized and integrated into
micro-systems, including parts for signal processing, providing great deal of advantages over
optical detection (Templin, Stoll et al. 2002). Because electrochemical reactions give an
electronic signal directly, there is no need for expensive signal transduction equipment.
Moreover, because immobilized biological sensing elements can be readily confined to a variety
of electrode substrates, detection can be accomplished with an inexpensive electrochemical
analyzer. Indeed, portable systems for clinical testing and on-site environmental monitoring are
now being developed (Wang 2002). So in this study, an electrochemical method is employed as a
transducer element in the biosensor design.
The fundamental process in an electrochemical method is the transfer of an electron between
the electrode surface and the molecules in the solution adjacent to the electrode. An
electrochemical detection system is composed of 2 major parts: 1) a potentiostat, which provides
potential or current and collects the electrical signal; 2) an electrochemical cell, which consists of
a certain volume of electrolytes and 3 electrodes, namely working, reference, and auxiliary
electrodes.
Figure 1.2 shows a schematic of a three-electrode electrochemical detection system (Bard
and Faulkner 2001). The working electrode is the one at which the reaction of electron transfer
occurs; the reference electrode (commonly silver/silver chloride) provides a stable and wellknown electrode potential against the working electrode; and the auxiliary electrode, coupled
with working electrode, facilitates the electron transfer in the circuit.

8

-

+

i
Auxiliary
electrode
Reference
electrode
Working
electrode

Figure 1.2 Schematic of a three-electrode electrochemical detection system. The system is
composed of a potentiostat, working, reference, and auxiliary electrodes. (adapted from Bard
and Faulkner 2001).
Electrochemical biosensors measure an electrochemical response of a biological event
directly or indirectly, of which there are four basic types: 1) conductometric, which is a change
in conductance of a bacterial cell between a pair of electrodes due to cell metabolism, such as
conductive polymer (Pal, Alocilja et al. 2007); 2) potentiometric, which is the difference in
electrical potential between a sample and a reference electrode (Karakus, PekyardImcI et al.
2006); 3) amperometric, which is a response due to oxidation or reduction of a specific chemical
at a constant applied potential (Hernandez-Santos, Diaz-Gonzalez et al. 2004; Karslan, Kayahan
et al. 2009); 4) Impedimetric, which is the change in the capacitance layer between electrodes
after biological binding (Kim, Park et al. 2009). All these electrochemical detection approaches
are typically fast, sensitive and low-cost.
Based on different sensing elements, the biosensor can be divided into immunosensor, DNA
sensor, cell-based biosensor, aptasensor, enzyme-based sensor, and other combinations. DNA
9

contains the genetic instructions used in the development and functioning of all known living
organisms and some viruses. Chemically, DNA consists of two long polymers of simple units
called nucleotides, with backbones made of sugars and phosphate groups. Attached to each sugar
is one of four types of molecules called bases. It is the sequence of these four bases along the
backbone that encodes information. Each type of base on one strand forms a bond with just one
type of base on the other strand. This is called complementary base pairing. Purines form
hydrogen bonds to pyrimidines, with adenine (A) bonding only to thymine (T), and cytosine (C)
bonding only to guanine (G). This arrangement of two nucleotides binding together across the
double helix is called a base pair.

Adenine Thymine

Guanine Cytosine

Sugar phosphate
backbone

Figure 1.3. Schematic of DNA base pairs.

10

Figure 1.3 shows a schematic of DNA. The base pair coding mechanism make the DNA
probe (pDNA) the most specific biological recognition molecules. The use of pDNA for isolating
and identifying gene sequences through hybridization is the most common in DNA sensor
research. Generally, a DNA sensor usually relies on the immobilization of a single-strand pDNA
onto a surface to recognize its complementary DNA target (tDNA) sequence by hybridization.
After specific hybridization between pDNA and tDNA, the transducer can convert the
hybridization to an electrical signal. In recent years, the interest for DNA-based diagnostic tests
has been growing. The development of systems allowing DNA detection is motivated by
applications in many fields: DNA diagnostics, gene analysis, fast detection of biological warfare
agents, and forensic applications (Sassolas, Leca-Bouvier et al. 2008). Moreover, these DNA
sensors are able to distinguish closely related species which most antibody tests could not, which
generate several other applications in the food industry. For example, with many countries
refusing to accept the import of genetically modified organism (GMO) food products, there is an
additional economic impact to their use. DNA sensors can be used to identify GMO products by
detecting transgenic genes (Kalogianni, Koraki et al. 2006; Sun, Zhong et al. 2008). DNA
biosensors have tremendous promise for obtaining sequence-specific information in a faster,
simpler and more cost-effective manner compared to the traditional molecular techniques.

1.5 Novelty of Research
The multiplex DNA sensor developed here is the first design of nanoparticle-based
electrochemical bio-barcode DNA detection based on a SPCE biosensor. The novelty is in the
PCR-less DNA recognition coupled with rapid, multiplex detection of pathogens on a disposable
SPCE biosensor. The bio-barcode DNA assay utilizes nanoparticle tracer (NT) terminated

11

oligonucleotides on gold nanoparticles (AuNPs) for signal amplification and magnetic
nanoparticles (MNPs) for easy and clean separation from the sample. The detection system does
21

not need PCR amplification due to its zeptomolar (10-

M) sensitivity (Nam, Stoeva et al. 2004).

The multiplex nanoparticle-based electrochemical bio-barcode DNA detection system provides a
new, simple, fast, and field-ready technology as an alternative to the traditional PCR technique
with comparable sensitivity. The biosensor design is very cost-effective to fabricate, is
environmental-friendly when disposed of, has rapid detection time, no surface modification on
the electrode surface, and is easy-to-miniaturize and integrate into micro-systems, and highly
portable for field applications -- all these parameters lending well as an alternative device for
bioterrorism and food safety countermeasure for prevention, protection, and mitigation.

1.6 Hypothesis
A nanoparticle-based electrochemical bio-barcode DNA sensor can simultaneously detect the
protective antigen A (pagA) gene (accession number = M22589) from Bacillus anthracis and the
insertion element (Iel) gene (accession number = Z83734) from Salmonella Enteritidis in the
same sample.

1.7 Specific Aims
To demonstrate proof of concept, the specific aims of the research are:
a) Synthesize and characterize gold nanoparticles and nanoparticle tracers (PbS and CdS);
b) Functionalize gold nanoparticles and magnetic nanoparticles, and conjugate biobarcoded AuNPs with nanoparticle tracers (NTs);
c) Evaluate and optimize SPCE chips for NT ion detection;

12

d) Confirm bio-barcode assay;
e) Evaluate biosensor sensitivity using a single purified PCR product from target organisms;
f)

Design multiplex detection of multiple targets in the same sample and evaluate its
sensitivity and specificity.

13

CHAPTER 2: LITERATURE REVIEW

2.1 Overview of Nano-sensors
Nanotechnology is the engineering of functional systems at an atomic and molecular scale.
The dimensional range of interest is approximately 1-100 nm. Because of effects such as the
quantum size effect, mini size effect, surface effect and macro-quantum tunnel effect,
nanomaterials or matrices with at least one of their dimensions ranging in scale from 1 to 100 nm,
display unique physical and chemical features. Through the different physico-chemical
phenomena taking place at nanodimensions, the integration of nanotechnology approaches into
biosensors holds great promise for addressing the analytical needs of pathogen detection. Novel
functional nanomaterials, such as porous silicon, nanowire, carbon nanotubes, and nanoparticles,
biosensors are exerting a profound influence on food safety, environmental monitoring, and
medical treatment (Staufer et al., 2007). Zhang and Alocilja (2008) reported a nano-scale porous
silicon (PS)-based biosensor for DNA detection. Due to a lot of nano-scale pore formation on the
silicon biosensor, much larger active surface area for hybridization was provided than planar
silicon, which leaded to an increase in signal. A bigger electrochemical response was observed
when hybridization occurred. When the concentration of tDNA increased, the charge transfer
between the redox marker and porous silicon electrode was enforced so that the peak current rose
with increasing DNA concentrations. The experiment result showed that the detection limit of
the PS-based label-free DNA biosensor was 1 ng/mL (Zhang and Alocilja 2008). A nanowire
labeled direct-charge transfer biosensor for detecting Bacillus species was reported (Pal, Alocilja
et al. 2007). Testing of the biosensor showed that it had the ability to detect the presence of the
1

target microorganism, Bacillus cereus, at concentrations as low as 10 CFU/ml, the detection

14

time being only 6 min (Pal, Alocilja et al. 2007). Carbon nanotubes, the essential element of
nanotechnology, have received considerable attention because of their inert properties,
conducting behavior, and high-surface area. Particularly, their promotional ability for electrontransfer reactions with enzymes and other biomolecules has made carbon nanotubes the ideal
supporting material for heterogeneous catalysts (Jianrong, Yuqing et al. 2004). A carbon multiwalled nanotubes (MWCNTs) based biosensor for direct and label-free detection of DNA of
influenza virus (type A) was reported (Tam, Van Hieu et al. 2009). The hybridization was
detected by changes in the conductance on the surface of sensors. The results showed that the
DNA sensor could detect as low as 0.5 nM of the target DNA samples; the response time of
DNA sensor was approximately 4 min (Tam, Van Hieu et al. 2009).
Besides porous silicon, conductive polymer, and carbon nanotubes, various nanoparticles
have raised great expectations with respect to generating enhanced signal-to-noise ratios,
reducing response times and using them in multiplexed systems (Khanna 2008). Nanoparticles
play most important roles in the development of the nano biosensor because: 1) Nanoparticles
exhibit higher ratios of surface area to volume than their bulk counterparts, so nanoparticle
modified electrochemical interfaces will provide larger electrochemically active areas and
therefore probably lead to higher detection sensitivity for target molecules; 2) some novel
nanoparticles, particularly metal nanoparticles, can easily act as enhancing agents for effective
acceleration of electron transfer between electrode and detection molecules, so leading to more
rapid current response for target molecules; 3) nanoparticles can act as a supramolecular
assembling unit with advanced functional properties for constructing a variety of architectures on
the surface of electrodes and further tailoring of an electrochemical-sensing interface; 4)
nanoparticles can be conjugated with some important biomolecules (e.g., redox enzyme) and also

15

act as nano-connectors that activate redox enzymes or electrical labels for bio-recognition events;
and, 5) nanoparticle-modified electrochemical interfaces behave as nanoelectrode ensembles
(Guo and Dong 2009). Inspired by these important features, literature already shows numerous
examples of incorporating nanoparticles into biosensors. So far, there are nano-biosensors for the
specific detection of biologically-relevant molecules (e.g., nucleic acids (Ting, Zhang et al. 2009;
Zhang, Carr et al. 2009), proteins (Lin, Qu et al. 2007) and enzymes (Ren, Meng et al. 2005) and
for the detection of infectious agents (Pal and Alocilja 2009).
Gold Nanoparticles (AuNPs)
Nanoparticles of noble metals, especially gold nanoparticles (AuNPs), have received great
interest due to their attractive electronic, optical, and thermal properties as well as catalytic
properties and potential applications in the fields of physics, chemistry, biology, medicine, and
material science and their different interdisciplinary fields (Pingarr, Yez-Sede et al. 2008). In
biosensor research, more attention has been paid to AuNPs because of their good biological
compatibility, excellent conducting capability and high surface-to-volume ratio. The unique
properties of gold nanoparticles to provide a suitable microenvironment for biomolecules
immobilization retaining their biological activity, and to facilitate electron transfer between the
immobilized proteins and electrode surfaces, have led to an intensive use of this nanoparticle for
the construction of electrochemical biosensors with enhanced analytical performance with
respect to other biosensor designs. Recent advances in the development of DNA biosensors using
gold nanoparticles to improve DNA immobilization on electrode surfaces and as suitable labels
to improve detection of hybridization events are considered.
Figure 2.1 shows the most important strategy used to integrate AuNPs in DNA-detection
systems (Castada, Alegret et al. 2007), comprising: (A) the electrochemical detection of AuNP

16

A

B

C

D

E

F

Figure 2.1. The different strategies used for the integration of gold nanoparticles (AuNPs) into
DNA-sensing systems: (A) previously dissolving AuNPs by using HBr/Br2 mixture followed by
Au(III)-ion detection; (B) direct detection of AuNPs anchored onto the surface of the genosensor;
(C) conductometric detection; (D) enhancement with Ag or Au followed by detection; (E)
AuNPs as carriers of other AuNPs; and, (F) AuNPs as carriers of other electroactive labels
(adapted from Castada, Alegret et al. 2007).
labels by detecting the gold ions released after dissolving them in acid; (B) direct detection of
AuNPs anchored onto the surface of a conventional genosensor (based on stripping voltammetry);
(C) silver enhancement using a conductometric technique; (D) enhancement of AuNPs anchored
to conventional genosensor surface by using silver or gold; (E) AuNPs as carriers of other
nanoparticles; and,(F) using AuNPs as carriers for other electroactive labels .
Table 2.1 summarizes some of the results obtained by using different strategies. Although
clear improvements have been demonstrated by the same authors upon comparing their results
(with and without enhancement) it is not so clear the improvement when comparing different

17

Table 2.1. Electrochemical genosensors using AuNPs as label. GE: gold electrode; DPV:
differential pulse voltammetry; ASV: anodic stripping voltammetry; SPMBE: sandwich-type
screen-printed microband electrode; PSA: potentiometric stripping analysis; SPEs: screenprinted electrodes; M-GECE: magnetic graphite-epoxy composite electrode, GCE: glassy carbon
electrode (Castada, Alegret et al. 2007).
AnNPs size Electrode
1.4 nm
20 nm
5 nm
5 nm
10 nm
5 ± 1.3 nm
16.3 nm
15 nm

M-GECE
SPMBE
SPEs
SPE
M-GECE
PGE
Chitosanmodified
GCE
SPMBE

Detection
Technique
DPV
ASV
PSA
PSA
DPV
DPV

Detection
mode
Direct
HBr/Br2
HBr/Br2
Direct
Direct
Direct

None
None
None
Silver
None
None

Detection
limits
12 nM
5 pM
5 ng
1.2 fmol
33 pmol
0.78 fmol

DPV

-

Silver

50 pM

ASV

HBr/Br2

Gold

600 aM

Enhancement

laboratories.
In this study, the good biological compatibility and high surface-to-volume ratio of AuNPs
were employed for biosensor design. AuNPs also were measured electrochemically as an
indictor of hybridization.
Magnetic Nanoparticle or beads (MNPs)
The biosensor detection is generally divided into two steps. The first is the capture, in which
the microbial species or group of interest, such as antigen and target DNA, is removed, tagged or
amplified to differentiate it from the remaining material in the sample. This step is typically
responsible for the selectivity of the approach. The second step is the detection, in which the
captured, tagged or amplified material is counted or measured quantitatively. The detector
typically acts as a transducer, translating the biological, physical, or chemical alteration into a
measurable electronic signal.

18

The current biggest technical challenges to the implementation of pathogen detection are the
sample processing and detection sensitivity. There are two possible approaches to overcoming
inadequate sensitivity: use the most advanced detectors available and more sensitive detector
technology, or preconcentration, which can enhance sensitivity several fold by increasing the
number of targets per unit volume at a relatively modest cost. As nanobiotechnology progresses,
sensors to detect pathogens or their constituents become smaller and more sensitive. Owing to
the nature of these nanoscale sensors, the sample size from which the detection is being made is
typically several microliter or smaller. Therefore, the main challenge for scientists developing
detection methods for pathogens is in the sample preparation now. Although the sample
preparation requirements will vary from one food product to another, research into this step is
required to bridge the emerging field of nano-biosensors with the food industry. The sample
preparation will not only depend on the food matrix, but on the pathogen as well. Pathogenic
viruses, bacteria and parasites might all exist within the same food product. For now, automation
using more traditional detection methods is increasing in popularity, and the same will be true for
less expensive and yet more powerful sensors of the future that are integrated with sample
preparation.
Since more detection technologies are based on measuring sample volumes less than 1 ml,
preconcentration may be necessary to achieve acceptable sensitivity. Several available modes of
preconcentration are being used, including filtration, size-fractionation, centrifugation and
magnetic separation or combinations of these methods. Magnetic nanoparticles (MNPs) or
magnetic beads (MBs) are frequently used for concentration, separation, purification and

19

Cells

Y

Specific antibody

antibody Y

Biotinylated
ss ODN

Y

Streptavidin
Protein G
protein A

antigen

Y
Oligo(dT)25

Complementary
NA strand

Cobalt chelate
His-tagged
protein

NA strand with
(A)n stretch
Figure 2.2. Magnetic beads (MBs) as versatile tools for bioassays. Beads modified with various
recognition elements (examples of commercially available MBs shown on the gray background)
can be used for specific bioaffinity capture of different molecules. For example, beads bearing
covalently attached oligo(dT)25 chains can bind nucleic acid involving (A)n stretches, including
natural eukaryotic mRNAs or tDNAs tagged with (A)n adaptors. Streptavidin-coated beads are
suitable for capturing any biotinylated molecules, including single strand or double strand
nucleic acids, aptamers, peptides, proteins, etc. Antibodies can be attached to the beads either via
a direct covalent linkage, or via specific antibody binding proteins such as protein A or protein G
(beads functionalized with the latter proteins can be further modified with various antibodies on
demand). Other affinity ligands attached to MBs are suitable for specific capture of tagged
recombinant proteins (such as metal-affinity cobalt chelate for histidine tags). Biomolecules
captured at the MB surface can serve as a biorecognition layer for interaction with other
molecules. For example, a biotinylated single strand oligonucleotide (ssODN) immobilized at
streptavidin-coated beads features a hybridization probe for complementary sequence, the
dsDNA can represent a target for a DNA-binding protein, antibody can bind a specific antigen

20

(including antigens exposed at surfaces of whole cells), immobilized protein (e.g., via antibody,
his-tagor biotin–streptavidin linkage) can interact with a specific nucleic acid or another protein,
etc. (adapted from Palecek and Fojta 2007).
identification of molecules and specific cells. They are versatile tools in the separation of nucleic
acids, proteins and other biomacromolecules, their complexes and cells. Figure 2.2 shows a
schematic of magnetic beads as versatile tools for bioassays (Palecek and Fojta 2007).
Magnetic methods rely on efficient separation of paramagnetic or ferromagnetic particles
from biological or chemical media. Superparamagnetic particles become magnetic under a strong
magnetic field but retain no residual magnetism in the absence of magnetic field. These particles
do not interact with each other when removed from the magnetic field. The diameter of the
particles is usually between 0.5 and 10 um. MNPs with hydrophilic surfaces are particularly
convenient for DNA oligonucleotides, providing reproducible magnetic separation. MNPs can be
prepared in various ways. Usually particles susceptible to magnetism, such as iron oxide, are
coated with biological or synthetic polymers.
A number of companies offer MNPs and kits optimally adjusted for the desired application.
Magnetic separators enabling automated handling of the beads are commercially available
(Berensmeier 2006). With the use of such systems a rapid diagnosis of infections (Satoh, Iwata et
al. 2003; Lee, Lien et al. 2008) , and extraction of DNA and RNA from various media (Hourfar,
Michelsen et al. 2005) is possible. New magnetic particles with improved properties, suitable for
diagnostics in microbiology, cell isolation and other purposes were developed. Recently a list of
12 companies and their products was published (Berensmeier 2006), and it can be expected that
materials with more specific binding properties and better separation will soon be developed.
The scale-up procedures for the purification of large volumes of biological agents and

21

miniaturization for sensors and lab-on-a-chip techniques can be foreseen. In this study, magnetic
particles were coated with DNA probe (pDNA) for sample separation and preconcentration.
Nanoparticle tracers (NTs)
Inorganic-colloid (quantum dots) nanoparticle tracers (NTs) are widely tagged to biological
molecules for proteomic function based-medical studies in post-genomic era (Bruchez, Moronne
et al. 1998), because NTs are dimensionally similar to biomolecules and a variety of
bioconjugates are ideally suited as optical or molecular imaging (Chan and Nie 1998). Quantum
confinement effects give rise to unique optical and electronic properties in quantum dots (QDs),
giving them numerous advantages over current fluorophores, such as organic dyes, fluorescent
proteins and lanthanide chelates. Properties that particularly influence fluorophore behavior, and
therefore applicability to different situations, include the width of the excitation spectrum, the
width of the emission spectrum, photostability, and the decay lifetime. So most current efforts
for multiplex assays have focused on multi-color fluorescent detection in connection with
different QDs. QDs have been used for the simultaneous fluorescent immunoassay of four toxins
(Goldman, Clapp et al. 2004). Goldman et al. (2004) used QDs functionalized with antibodies to
perform multiplexed fluoroimmunoassays for simultaneous detection of various toxins. This
sensor could be used for environmental purposes for concurrently recognizing pathogens like
cholera toxin in water. The fluorescence resonance energy transfer analysis (FRET) principle in
QDs was also applied to a maltose biosensor (Medintz and Deschamps 2006). The sensing
mechanism was the application of semiconductor QDs conjugated to a maltose binding protein
covalently bound to a FRET acceptor dye. In absence of maltose, the dye occupied the protein
binding sites. Energy transference from the QDs to the dyes quenched the QD fluorescence.
When maltose was present, it replaced the dye leading to recovery of the fluorescence.

22

Compared to previous results using organic fluorophores, QDs gave an increased luminescence
of 52% over 47 h after incubation with a collagenase (Medintz and Deschamps 2006).
Though current optical sensing approaches using QDs are effective for high-density arrays,
they are often in the laboratory, require complicated equipment and special training, and
complicated by the requirement of an elaborate excitation and detection scheme and by the broad
emission bands so that they are hard to be miniaturized and portable. While electrochemical
systems for biomedical diagnostics can be miniaturized and integrated into systems, including
parts for signal processing, to provide great deal of advantages over optical detection schemes.
They are not as susceptible to turbidity interference as optical-based detection, and typically have
very low detection limits. So they are ideally suited for meeting the portability requirements of
on-field detection of pathogens. QDs used for the electrochemical hybridization detection of
multiple DNA targets was reported (Liu, Wang et al. 2004). Different inorganic quantum dots
(QDs), namely nanoparticles tracers, (ZnS, CdS, and PbS) tagged to DNA or antibody for
biological molecules detection are reported (Liu, Wang et al. 2004). The biosensor could detect
multiplex proteins in a mixture containing 100 ng/mL β2-microglobulin, IgG, BSA, and CRP in
connection with ZnS-, CdS-, PbS-, and CuS-labeled antibodies, respectively. Baseline-resolved
peaks of nearly similar sensitivity and favorable signal-to-noise characteristics were observed for
the four proteins. Their metal components yielded well resolved highly sensitive stripping
voltammetric signals for the corresponding targets (Liu, Wang et al. 2004). Nanoparticle tracers
also were used to detect DNA hybridization based on the use of different nanoparticle tracer tags
(Wang, Liu et al. 2003). This approach could be multiplexed and scaled by incorporating
additional quantum-dot tracers (including bimetallic ones), by using microtiter plate platforms

23

(with each microwell carrying out multiple measurements), and through particle-based libraries
for electrical barcoding in the future.

2.2 Electrochemical DNA sensor
Successful recognition of a specific sequence of DNA requires a highly specific recognition
layer. DNA biosensors exploit the ability of a single strand of DNA (ssDNA) to recognize and
hybridize with a complementary ssDNA in the recognition interface. The strand of DNA, which
does the detection, the DNA probe, is immobilized onto the surface of the signal transducer (see
Figure 2.3). Exposure of the recognition surface to a sample containing single strands of the
DNA of interest, the target DNA, will result in hybridization where transduction can proceed.
In a DNA biosensor, the probe will most frequently consist of an oligonucleotide of about 15
to 50 bases long. To a large extent the selectivity, sensitivity and reproducibility of a DNA
recognition interface will depend on the immobilization of the DNA probe (pDNA). When
hybridization occurs, the probe and target strands must be free to coil around each other.
Immobilization of the probe strand onto a transducer surface will inevitably cause a decrease in
configurational freedom. Therefore minimization of the decrease in this freedom is required to
achieve efficient hybridization. Single point attachment of the probe strand at either the 5’ or 3’
end of a pDNA is one strategy to maintaining efficient hybridization. Alkanethiols are molecules
with an alkyl chain, (C-C)n chain, as the back bone, a tail group, and a SH head group. They are
used on noble metal substrates because of the strong affinity of sulfur for these metals and form a
self assembled monolayer (SAM) on it. In this study, thiolated DNA probes (pDNA) and barcode
DNA (bDNA) were coated on AuNPs homogeneously to improve the hybridization efficiency of
pDNA and tDNA.

24

DNA target

Probe

A
G
T
C
A
G

A
G
T
C
A
G

Hybridization

Transducer

.
T
G
A
=T
=C
=A
=G
=T
=C

Signal
Processing

Transducer

(A)

(B)

(C)

Figure 2.3. Schematic of an electrochemical DNA biosensor. (A) DNA probe immobilization; (B)
hybridization between DNA probe and target DNA on the transducer; (C) signal processing by a
biosensor system.
The basic categories of electrochemical detection of DNA include direct reduction/oxidation
of DNA (Karadeniz, Gulmez et al. 2003; Ozkan-Ariksoysal, Tezcanli et al. 2008), indirect
oxidation of tDNA through the use of electrochemical mediators (Yang and Thorp 2001), DNAspecific redox indicator detection (Miroslav Fojta 2003), DNA-mediated charge transport
electrochemistry (Steel, Herne et al. 1998; Jackson and Hill 2001), and nanoparticle-based
electrochemical amplification (Sun, Zhong et al. 2008). Table 2.2 shows comparison of
platforms for DNA electrochemical sensing (Drummond, Hill et al. 2003).
Due to high background current, the direct detection of guanine (G) or adenine (A) on a
DNA sequence is still not sensitive and not ideal for multiplex detection. Methods to oxidize
tDNA indirectly through the use of electrochemical mediators have also been explored (Yang
2+

and Thorp 2001). An especially attractive approach uses polypyridyl complexes of Ru
Os

2+

and

to mediate the electrochemical oxidation of guanine. DNA- specific redox indicator

detection is an analogy to fluorescence-based methods.

25

Table 2.2 Comparison of Platforms for DNA electrochemical sensing (Drummond, Hill et al.
2003).
Type of sensor
Direct DNA
electrochemistry
Indirect DNA
electrochemistry

Advantages
Highly sensitive (femtomoles of target);
requires no labeling step; amenable to
range of electrode
Highly sensitive (attomoles of target);
usually requires no labeling step;
multiple-target detection at same electrode

DNA-specific
redox indicator
detection

Moderate to high sensitivity (femtomoles
of target); well suited to multiple-target
detection; samples remain unaltered

Nanoparticlebased
electrochemistry
amplification

Extremely sensitive (femtomole to
zeptomole range); well suited to multipletarget detection with different
nanoparticles

DNA-mediated
charge transport

Highly sensitive (femtomole range) and
simple assay; requires no labeling;
uniquely well suited for mismatch
detection; sequence independent;
amenable to multiplexing; applicable to
DNA-protein sensing step

The reporting

pDNAs

are

labeled

with

Disadvantages
High background signals;
can not be multiplexed;
destroy the sample
Probe substrate can be
difficult to prepare; destroys
the sample
Chemical labeling step
required unless “sandwich”
method used; sequence
variations can be problematic
Many development steps in
assay; reliability and
robustness of surface
structure problematic;
sample usually destroyed
Biochemical preparation of
target sample required

redox-active molecules.

Appearance

of

the characteristic electrochemical response of the redox reporter therefore shows the
10

hybridization event. Detection limits on the order of 10

molecules have been reported

(Miroslav Fojta 2003). DNA-mediated charge transport electrochemistry uses a redox active
reporter molecule, such as intercalator, to bind the double helix of DNA noncovalently and
facilitate the charge transport through DNA structure. It is uniquely suited to sense changes in
DNA damage, mistakes, and mismatches.
Currently many applications of DNA sensing involve extremely small numbers of target
analytes, with correspondingly few hybridization events. Many nanomaterials were deposited

26

onto traditional electrodes to improve the biosensor sensitivity. Multiwalled carbon nanotubes
(MWNT) are receiving more and more attention interests in biosensor research. A MWNT
nanoelectrode arrays embedded in a silicon dioxide (SiO2) matrix for DNA detection was
reported (Jessica Koehne 2003). The inherent guanine bases in the DNA amplicon target of 300
2+

bases serve as signal moieties with the aid of tris(2,2‘-bipyridine) ruthenium (II) (Ru(bpy)3 ),
providing an amplified anodic current associated with the oxidation of guanine groups at the
nanoelectrode surface. The reduced size and density of the nanoelectrode array provided by
MWNTs dramatically improves the sensitivity of electrochemical detection. In addition, the
abundant guanine bases in tDNA produce a large signal. Less than 1000 target amplicons can be
detected on a microspot, approaching the sensitivity limit of conventional laser-based
fluorescence techniques. This method also eliminates the labeling requirement and makes the
measurements much simpler (Jessica Koehne 2003). It was reported that lead selenide
nanoparticle/chitosan composite films were prepared on the carbon paste electrode (CPE) for the
detection of CaMV35S promoter genes in some transgenic plants (Xie, Jiao et al. 2008). The
–11

result showed the detection range from 5.0 × 10
–11

× 10

–6

to 5.0 × 10

M and the detection limit of 1.6

M. using this DNA electrochemical sensor.

Analyte amplification has also been accomplished in an innovative way indirectly using
electrochemical detection of nanoparticles. Semi-conducting nanoparticle tracers (NPs) are good
oligonucleotide labeling markers for DNA biosensors. For example, an electrochemical coding
technology for the simultaneous detection of multiple DNA targets based on nanoparticle tracers
with different redox potentials was reported (Wang, Liu et al. 2003). Such encoding
nanoparticles thus offer a voltammetric signature with distinct electrical hybridization signals for

27

the corresponding DNA targets. The new electrochemical coding bioassay relies on the use of
different inorganic-colloid (quantum dots) nanoparticle tracers, whose metal components yield
well resolved highly sensitive stripping voltammetric signals for the corresponding targets. Three
encoding nanoparticles (zinc sulfide, cadmium sulfide, and lead sulfide) have thus been used to
differentiate the signals of three DNA targets in connection with a sandwich hybridization assay
and stripping voltammetry of the corresponding heavy metals.
Figure 2.3 shows the schematic of multi-target DNA detection using different inorganic
colloid nanoparticle tracers. The new multi-target electrical detection scheme incorporated the
high sensitivity and selectivity advantages of nanoparticle-based electrical assays. This approach
could be

multiplexed and scaled by incorporating additional nanoparticle tracers, by using

microtiter plate platforms (with each microwell carrying out multiple measurements), and

PbS

CdS

PbS

Zn

Pb

ZnS
Me
Me

CdS
A

Cd

2+

0

Me

2+

ZnS

B

C

D

Figure 2.4. Multi-target electrical DNA detection protocol based on different inorganic colloid
nanoparticle tracers. (A) Introduction of probe-modified magnetic beads. (B) Hybridization with
the DNA targets. (C) Second hybridization with the QD-labeled probes. (D) Dissolution of QDs
and electrochemical detection (adapted from Wang, Liu et al. 2003).

28

through particle-based libraries for electrical barcoding in the future. In addition, surface
modification of electrodes the immobilization of pDNA was not necessary. It will save a lot of
time of electrode preparation and prolong the shelf life of electrodes greatly.

2.3 Bio-barcode Assay
Most of the current DNA sensors need PCR amplification before applying the samples
because of relatively low sensitivity. However, PCR is often criticized for its complex, expensive,
time-consuming, and labor-intensive procedure. It limits the widespread application of DNA
sensor.
The bio-barcode DNA assay is a promising new amplification and detection technique that
makes use of short oligonucleotides as target identification strands and surrogate amplification
units in both protein and nucleic acid detection (Hill and Mirkin 2006). The technique uses many
advantageous properties of oligonucleotide-functionalized AuNPs including ease of fabrication,
greater oligonucleotide binding capabilities, stability under a variety of conditions, catalytic
ability, and optical properties (Demers, Ostblom et al. 2002; Lytton-Jean and Mirkin 2005). The
large ratio between thiolated single-strand oligonucleotide barcodes DNA (bDNA) and pDNA on
AuNPs provides significant amplification. It has been shown to have extraordinarily PCR-like
–18

sensitivity. Under controlled conditions, the assay has shown attomolar (10

–21

variety of protein targets (Nam, Thaxton et al. 2003) and zeptomolar (10

) sensitivity for a

) sensitivity for a

variety of target genes (Nam, Stoeva et al. 2004). The bio-barcode DNA assay utilizes
oligonucleotide-modified gold nanoparticles (AuNPs) for signal amplification and magnetic
nanoparticles (MNPs) for easy and clean separation from the sample. Figure 2.5 shows a
schematic of bio-barcode DNA assay. After immobilization of oligonucleotides on the surface of

29

nanoparticles, both nanoparticles can bind with the target DNA (tDNA) to form a sandwich
structure, due to the specificity of DNA probe (pDNA). Figure 2.5A shows the tDNA sample
hybridizing with the second pDNA
(2pDNA) on the MNPs, forming a MNP-2pDNA/tDNA complex (middle picture). Then AuNPs
coated with the first pDNA (1pDNA) and bDNA are added to form a sandwich structure
consisting of MNP-2pDNA/tDNA/1pDNA-AuNP-bDNA (picture on the right). After
hybridization is complete, the sandwich structure is separated magnetically from unreacted
pDNA-AuNP-bDNA. Finally the bDNA is released from the surface of AuNPs (Figure 2.5B).
Based on different labels, the barcode strands can be identified on a microarray via scanometric
A

tDNA

MNPs2pDNA

The composite of
MNPs-2pDNA
/tDNA

B

The composite of
MNPs-2pDNA
/tDNA/1pDNAAuNP-bDNA
bDNA
release

Probe
separation
bDNA
Magnetic field
Figure 2.5. Schematic of bio-barcode DNA assay. (A) formation of MNP-2pDNA
/tDNA/1pDNA-AuNP-bDNA sandwich structure; (B) probe separation and bDNA release.

30

detection (Taton, Mirkin et al. 2000; Stoeva, Lee et al. 2006), on-chip detection (Goluch, Nam et
al. 2006), fluorescence (Oh, Nam et al. 2006), or Raman active dye (Cao, Jin et al. 2002) if the
barcodes carry with them a detectable marker.
2.4 Screen-printed Carbon Electrode
Solid electrodes based on carbon materials are commonly used in electrochemical analysis
due to their broad potential window, low background current, rich surface chemistry, low cost,
chemical inertness, and suitability for various sensing and detection applications (Kissinger and
Heineman 1996). The screen-printing microfabrication technology is nowadays well established
for the mass production of thick film electrodes. This process implies the sequential deposition
of layers of different conductive (carbon) or non-conductive inks on a variety of inert substrates.
The screen-printed carbon electrode (SPCE) is made of electrochemically active graphite
particles. A binder is added to enhance the affinity of the ink to the substrate in terms of adhesion
properties and mechanical strength. Solvents are used to improve the viscosity of the ink for the
printing process. Compared with conventional electrodes, the SPCE has several advantages, such
as simplicity, convenience, low cost and the avoidance of contamination between samples
(Susmel, Guilbault et al. 2003; Miscoria, Desbrieres et al. 2006). It represents a promising route
for mass production of inexpensive, reproducible and disposable electrochemical sensors for the
determination of trace levels of important compounds. There has been a lot of published work on
the detection of metals with screen-printed carbon electrodes using stripping analysis techniques
(Desmond, Lane et al. 1998; Kadara and Tothill 2004; Hwang, Han et al. 2008). Biosensors
based on screen-printed carbon electrode (SPCE) have been extensively used for detection of
glucose (Guan, Li et al. 2005; Xu, Li et al. 2005), cholesterol (Carrara, Shumyantseva et al.
2008), antigens (Kim, Seo et al. 2006) and DNA (Ruffien, Dequaire et al. 2003; Hernandez-

31

Santos, Diaz-Gonzalez et al. 2004). However, these electrochemical detection systems with
electrochemical sensors need additional auxiliary and reference electrodes and a relatively large
volume of electrolyte solution. This limits potential applications of electrochemical sensors for
portable detection, especially for samples with a very small volume requirement such as body
fluids. A SPCE integrated with an auxiliary/reference electrode was used in this study. Its
volume requirement was 100 µl. The potential application for portable detection is promising.
The SPCE can be enhanced with mercury or bismuth. The mercury-film electrode (MFE) and
hanging mercury drop electrode (HMDE) are commonly used in the development of stripping
voltammetry. However, because of the toxicity of mercury, future regulations and occupational
health considerations may severely restrict its use as an electrode material. Bismuth is an
environmentally friendly element, with low toxicity and a widespread pharmaceutical use
(Rodilla, Miles et al. 1998). The bismuth electrodes can be prepared by simultaneous deposition
of bismuth and the target heavy metals, in a manner analogous to in situ plated mercury film
electrodes. Such electrodes display well-defined, sharp and highly reproducible stripping peaks
for low (ppb) concentrations of lead, cadmium, or zinc over a low background current. Such use
of "mercury-free" electrodes is particularly attractive for the development of multiplex metal
sensors (Wang, Lu et al. 2000). In this study, a disposable integrated SPCE chip enhanced by
bismuth was used for the multiplex detection of NT ions in 100 µL of sample solution, which
were released from bio-barcode assays.

2.5 Square wave anodic stripping voltammetry
Square wave anodic stripping voltammetry (SWASV) is a powerful tool for measuring trace
metal ions. It can be viewed as combining the best aspects of square wave voltammetry (SWV)

32

and anodic stripping voltammetry (ASV). The technique includes the background suppression
and sensitivity of differential pulse voltammetry, and the diagnostic value of anodic stripping
voltammetry (Bard and Faulkner 2000). In square wave voltammetry, a square wave is
superimposed on the potential staircase sweep. Oxidation or reduction of species is registered as
a peak or trough in the current signal at the potential at which the species begins to be oxidized
or reduced. Figure 2.6 shows waveform and measurement scheme for square wave voltammetry
(Bard and Faulkner 2000).
The current is measured at the end of each potential change, right before the next, so that the
contribution to the current signal from the capacitive charging current is minimized. The
differential current is then plotted as a function of potential, and the reduction or oxidation of
species is measured as a peak or trough. Due to the lesser contribution of capacitative charging

Potential

tp

Forward
sample

ΔEp
Reverse
sample

ΔEs

Time
Figure 2.6. Waveform and measurement scheme for square wave voltammetry. Shown in bold is
the actual potential waveform applied to the working electrode. The light intervening lines
indicate the underlying staircase onto which the square wave can be regarded as having been
superimposed. In each cycle, a forward current sample is taken at the time indicated by the solid
dot, and a reverse current sample is taken at the time marked by the shaded dot (adapted from
Bard and Faulkner 2000).

33

Deposition
Preelectrolysis

Rest period

Stripping

Stirrer
off
icath

+n

M M +ne
ts

+n

M +ne M
td

ianod

t

tr
ip

(a)

Ed

(b)

(c)

E

Ep
t

Figure 2.7. Principle of anodic stripping. Values shown are typical ones used; potentials and Ep
2+

are typical of Pb analysis, (a) Preelectrolysis at Ed; stirred solution, (b) Rest period; stirrer off.
(c) Anodic scan (v = 10-100 mV/s) (adapted from Bard and Faulkner 2000).
current the detection limits for SWV are on the order of nanomolar concentrations (Bard and
Faulkner 2000).
Besides the advanced measurement procedure of SWV that generates an extremely favorable
signal-to-background ratio, the remarkable sensitivity of SWASV technique is also attributed to
the combination of an effective pre-concentration step of anodic stripping techniques. Figure 2.7
shows the principle of anodic stripping. The analyte of interest is electroplated on the working
electrode during a deposition step, and oxidized from the electrode during the stripping step. The
current is measured during the stripping step. The oxidation of species is registered as a peak in
the current signal at the potential at which the species begins to be oxidized. The stripping step
can be either linear, staircase, square wave, or pulse (Bard and Faulkner 2000).
34

SWASV is a powerful electroanalytical technique for trace metal measurements. Because of
its effective “built-in” preconcentration (deposition) step, this technique offers remarkably low
(picomolar) detection limits. Four to six metals could be measured simultaneously in various
-10

matrixes at concentration levels down to 10

M, utilizing relatively inexpensive and portable

instrumentation (Wang 2003).
Recent activity has led to highly sensitive nanoparticle-based stripping electrical bioassays
based on capturing various colloidal metal and inorganic-crystal nanoparticle tags. Three metal
sulfide nanoparticle tracers (zinc sulfide, cadmium sulfide, and lead sulfide) have been used to
differentiate the signals of three DNA targets in connection with a sandwich hybridization assay
and stripping voltammetry of the corresponding heavy metals (Wang, Liu et al. 2003). This
electrochemical coding bioassay relied on the use of different inorganic-colloid nanoparticle
tracers (quantum dots) as electrochemical indicators, whose metal components yielded well
resolved highly sensitive stripping voltammetric signals for the corresponding targets. The new
strategy thus combined a novel multi-target bio-detection with an inherently amplified signal and
the high selectivity attribute of magnetic assays.

35

CHAPTER 3: MATERIALS AND METHODS

3.1 Reagents and Materials
Hydrogen tetrochloroaurate (III) trihydrate and sodium citrate dehydrate were used for the
synthesis of gold nanoparticles. Sodium sulfide, 3-mercaptoacetic acid, lead nitrate and cadmium
chloride were used for the synthesis of lead sulfide (PbS) and cadmium sulfide (CdS) NTs. 1,4Dithio-DL-threitol (DTT) was used for the cleavage of oxidized thiolated oligonucleotides. 1Ethyl-3-[3-dimethylaminopropyl]

carbodiimide

Hydrochloride

(EDC)

and

N-

hydroxysuccinimide (NHS) were used for the conjugation of carboxylic group on NTs and amine
group on bio-barcoded AuNPs. Amine terminated MNPs were used for sample separation. All
reagents were purchased from Sigma (St. Louis, MO). To validate and characterize SPCE chips,
potassium ferricyanide, potassium chloride, lead chloride and cadmium chloride were purchased
from Sigma (St. Louis, MO, USA). Bismuth standard stock solution (1000 mg/L, atomic
absorption standard solution) was obtained from Fisher Scientific (Hanover Park, IL, USA).
DNA isolation was performed using the QiaAmp DNA Mini Kit (Qiagen Inc., Valencia). Nap-5
column (GE Healthcare, Piscataway, NJ) was used to purify the DNA product from DTT
solution. Sulfosuccinimidyl 4-N-maleimidomethyl cyclohexane-1-carboxylate (sulfo-SMCC;
Pierce, WI) was used as a cross-linker between thiolated DNA probe (pDNA) and amine-coated
MNPs. Sulfo-NHS acetate (Pierce, WI) was used to block unreacted sulfo-SMCC. Bismuth
standard stock solution (1000 mg/L, atomic absorption standard solution) was obtained from
Fisher Scientific (Hanover Park, IL, USA). All solutions were prepared in deionized water (~ 18
MΩ cm).

36

3.2 Facilities and Equipment
Target DNA (tDNA) was amplified by a thermocycler (Mastercycler Personal, Eppendorf).
Gel electrophoresis (Runone System) was used to confirm the PCR product. After purification of
the PCR product, a spectrophotometer (SmartSpec 3000, Bio-Rad Laboratories) was used to
measure the concentration of pDNA in solution and tDNA in the sample. In the characterization
experiments of AuNPs, a UV-Vis-NIR Scanning Spectrophotometer (UV-3101PC, Shimadzu)
was used to determine the absorbance of AuNPs. A transmission electron microscope (TEM,
JEOL100 CXII) was used to image and characterize the NTs and AuNPs. All magnetic
separation was done using a magnetic separator (FlexiMag, SpheroTech). A centrifuge (Micro12,
Fisher Scientific) was used to separate and purify the nanoparticles. An incubator (HS-101,
Amerex Instrument Inc.) was used for the hybridization reaction. A fluorescent multi-label
counter (Victor 3, PerkinElmer) was used for measuring the fluorescent signal. Electrochemical
measurement was performed with a potentiostat/galvanostat (263A, Princeton Applied Research,
MA) that is connected to a personal computer. The electrochemical software operating system
(PowerSuite, Princeton Applied Research, MA) was used for electrochemical measurement and
data analysis. The integrated SPCE chips were purchased from Gwent Inc. England. Figure 1
shows a schematic of the integrated SPCE chip, which is composed of two electrodes: carbon
electrode (working electrode), and silver/silver chloride electrode (auxiliary and reference
electrode). The working area is limited by a meshed well. The volume capacity of the
electrochemical cell is 100 µL.

37

Working electrode
(carbon electrode)

A

B

Electrochemical cell
(insulating well)

Substrate
Auxiliary/reference
electrode (Ag/AgCl)

Figure 3.1. Schematic of screen-printed carbon electrode: A) top view, B) cross-section view. It
is composed of working electrode (carbon electrode) and auxiliary/reference electrode (Ag/AgCl)
in an insulating well.

3.3 Synthesis of Nanoparticles
Gold nanoparticles were synthesized by a chemical reduction method (Hill and Mirkin, 2006).
Hydrogen tetrochloroaurate (III) trihydrate aqueous solution (1 mM, 50 mL) was prepared in a
flask. The gold solution was heated while stirring on a hotplate. Once it refluxed vigorously, the
solution was slowly titrated with 5 mL of 38.8 mM sodium citrate. The solution turned from
yellow to clear, to black, to purple and finally deep red. The UV–VIS absorption spectrum of the
AuNPs solution was measured by a UV-Vis-NIR Scanning Spectrophotometer (UV-3101PC,
Shimadzu Inc.).
Nanoparticle tracers (PbS and CdS) were prepared according to the literature (Zhu, Zhang et
al. 2004; Ding, Zhang et al. 2009) by using 3-mercaptoacetic acid as a stabilizer. Briefly, 3mercaptoacetic acid (9.22 μL) was added to 50 mL of 0.4 mM Pb(NO3)2 solution; pH was
adjusted to 7 with 0.5 M NaOH. The solution was bubbled with nitrogen for 30 min, then 1.34
mM Na2S was dropwise added to the mixture. The reaction was carried out for 24 h under
bubbled nitrogen, and then gradually a brown colloid was formed. For CdS nanoparticles, 2 μL
of 3-mercaptoacetic acid was added to 100 mL of 1 mM CdCl2 solution; pH was adjusted to 11
using 0.5 M NaOH. The solution was bubbled with nitrogen for 30 min, then 50 mL of 1.34 mM
38

Na2S was dropwise added. The reaction was carried out for 24 h under bubbled nitrogen, and a
yellowish colloid was gradually formed. Both NTs were characterized using TEM.

3.4 Bacteria Culture, DNA isolation, PCR and DNA Probes
A clinical strain of Salmonella Enteritidis (strain S-64) and Bacillus anthracis Sterne strain
was used in this study. The pathogen was grown following standard microbiological cultures.
After enrichment, the cells were enumerated by spiral plating appropriately diluted cultures on
agar. DNA isolation was performed from 1 mL of culture using the QiaAmp DNA Mini Kit
(Qiagen, Valencia, CA).
The insertion element (Iel) gene (accession number Z83734) is chosen as the target of
Salmonella Enteritidis because the sequence’s specificity has been well studied in the literatures
(Wang and Yeh 2002; Song, Ahn et al. 2005). The pagA gene (accession number M22589) from
Bacillus anthracis Sterne strain will be used as the second target gene because Bacillus anthracis
virulent strain and Bacillus anthracis Sterne strain have this genotypic similarity (Song, Ahn et
al. 2005).
Primers and pDNAs for the insertion element (Iel) gene of Salmonella Enteritidis were as
follows.

The

single

stranded

forward

and

reverse

primers

were

IelL-5’-

CTAACAGGCGCATACGATCTGACA-3’ and IelR-5’-TACGCATAGCGATCT CCTTCGTT
G-3’. The following thiolated oligonucleotides were used to conjugate with the nanoparticles:
The first pDNA (1pDNA) on AuNPs: 5’-AATATGCTGCCTACTGCCCTACGCTT-SH-3’; the
second pDNA (2pDNA) on MNPs: 5’-SH- TTTATGTAGTCCTGTATCTTCGCCGT-3’ (Wang
and Yeh 2002); bDNA on AuNPs: 5’-TTATTCGTAGCTAAAA AAAAAA-SH-3’(Hill and
Mirkin 2006).

39

The primers and pDNAs of the pagA gene (accession number = M22589) from Bacillus
anthracis Sterne strain were as follows: pagAL: 5´-AAAATGGAAGAGTGAGGGTG-3´;
pagAR: 5´-CCGCCTTTCTACCAGATTTA-3´; 1pDNA: 5´-SH-GGAAGAGTGAGGGTGGAT
ACAGGCT-3´; 2pDNA: 5´-AGATTTAAATCTGGTAGAAAGGCGG–SH-3´ (Song, Ahn et al.
2005; Hill and Mirkin 2006).
TEX 613 (λexcitation = 596 nm, λemission = 613 nm) was used to label the bDNA and 6 Carboxyfluorescein (6-FAM; λexcitation = 495 nm, λemission = 520 nm) was used to label the
2pDNA on MNPs. Both TEX 613 and 6-FAM were used to evaluate the conjugation efficiency.
All oligonucleotides were synthesized by the Integrated DNA Technologies Inc. (Coralville, IA).
The PCR products were identified by gel electrophoresis and a UV transilluminator after
purification. After PCR amplification, the PCR product was purified by PCR purification kits
(QIAquick; QIAgen, Valencia, CA) and diluted serially with distilled water. PCR product
concentration and quality were measured with a Bio-Rad SmartSpec 3000 spectrophotometer.
The serially diluted PCR products of tDNA were used as DNA samples for the experiments.

3.5 Functionalization of Nanoparticles
To ensure full reactivity, thiol-modified oligonucleotides should be reduced immediately
before use. Otherwise, the thiol group on the oligonucleotides could not form a self-assembled
monolayer on the surface of AuNPs due to loss of active thiol group. DTT solution (0.1 M) was
prepared in the disulfide cleavage buffer (170 mM PBS buffer, pH=8.0). The thiolated
oligonucleotide was suspended in 100 µL DTT solution and the solution was allowed to stand at
room temperature for 2 hours. Nap-5 columns were then used for purifying the reduced thiolated
oligonucleotides. The procedure was followed according to the manufacture’s manual. After

40

purification, a UV-visible spectrophotometer was used to determine the DNA concentration. The
AuNPs synthesized previously (1mL), the purified thiolated bDNA (5 nmol), and the purified
thiolated 1pDNA (0.05 nmol) were then mixed together. Thiolated DNA would form a selfassembled monolayer on the surface of AuNPs. Figure 3.2 shows a schematic of the AuNPs
functionalization. After a serial salt addition (Hill and Mirkin, 2006), the particles were stabilized
for long-time storage at room temperature.
For the MNPs, the polyamine-functionalized iron oxide particles (1 mg) were reacted with
300 µg of sulfo-SMCC bifunctional linker for 2 h in 1 mL coupling buffer (0.1M PBS buffer,
0.2M NaCl, pH = 7.2). The supernatant was removed after magnetic separation and the
MNP-cross-linker conjugate were rinsed with the coupling buffer 3 times. The reduced thiolated
2pDNA (1 nmol) was added into 1 mL coupling buffer containing sulfo-SMCC-modified MNPs
and reacted for 8 h.

DNA functionalization

Gold
Nanoparticle
Thiolated bDNA
Thiolated 1pDNA

The composite of 1pDNAAuNP-bDNA

Figure 3.2. Schematic of the functionalization of AuNPs. A mixture of thiolated oligonucleotides
(bDNA:1pDNA=100:1) form a self assembled monolayer on the surface of gold nanoparticles.

41

MNPs -NH

MNPs -NH

HS-

Thiolated 2pDNA

Maleimide-activated
MNPs

MNPs
Sulfo-SMCC

-S-NH

MNP-2pDNA conjugate

Figure 3.3. Schematic of the functionalization of MNPs. (1) formation of maleimide-activated
MNPs; (2) formation of MNP-2pDNA conjugate.
Figure 3.3 shows the schematic of the MNP-pDNA conjugation. After conjugation, the
pDNA-immobilized MNPs were rinsed with the coupling buffer 3 times. The unreacted pDNA in
the supernatant was collected for the evaluation of conjugation efficiency. The functionalized
MNPs were then suspended in 35 mL of 10 mM sulfo-NHS acetate. The solution was incubated
and shaken at room temperature to block the unreacted sulfo-SMCC on the surface of MNPs.
After passivation, the particles were centrifuged at 1100 g for 1 min and washed with passivation
buffer (0.2M Tris, pH=8.5) and then with a storage buffer (10 mM PBS buffer, 0.2M NaCl,
pH=7.4).
Fluorescein-labeled 2pDNA and bDNA were used to evaluate the conjugation efficiency. To
separate the conjugates from unreacted oligonucleotides, different separation procedures were
taken. After centrifuging at 12,000 g for 20 min, the fluorescence signal of the supernatant
solution of 1pDNA/AuNPs/bDNA-TEX 613 was measured. The supernatant solution of
MNPs/Sulf-SMCC/2pDNA-6-FAM was measured after magnetic separation. The same

42

separation procedure was applied to their respective controls. Figure 3.4 shows the evaluation of
conjugation efficiency: MNP- pDNA and AuNP-bDNA.
To conjugate NTs with bDNA on AuNPs, N-hydroxysuccinimide (NHS) and 1-Ethyl-3-[3dimethylaminopropyl] carbodiimide Hydrochloride (EDC) were used to link the carboxylic
group on NTs and amine group on bDNA. EDC (5 mg) was added to 25 μL NTs, then 50 μL of
9% NHS in dimethyl sulfoxide were added. Hydrochloride ( 775 μL, pH 5) was added
immediately. The reaction was carried out on a vortex mixer at room temperature for 20 min. To
keep the optimal pH value of the reaction, 5 µL of phosphate buffer (1 M NaCO3, pH 8) was
added. The solution was shaken for 8 hours at room temperature and the conjugation solution
was washed with water at 12,000 g 3 times before usage.

Sample:
Fluorescein
-labeled
DNA probe

Centrifugation
or magnetic
separation

Fluorescein
-labeled
DNA probe

Centrifugation
or magnetic
separation

Control:

Figure 3.4 Evaluation of conjugation efficiency: MNP- pDNA and AuNP-bDNA.

43

Functionlizaton

EDC+NHS

AuNP
Thiolated bDNA with amine group
Thiolated 1pDNA

1pDNA-AuNP-bDNANTs complex

NT with carboxylic group
Figure 3.5. Schematic of conjugation of bio-barcoded AuNP with NTs. (1) functionalization of
gold nanoparticles with thiolated bDNA and thiolated 1pDNA; (2) formation of the 1pDNAAuNP-bDNA-NTs complex by cross-linking amine group on bDNA and carboxylic group on
NTs though 1-Ethyl-3-[3-dimethylaminopropyl] carbodiimide Hydrochloride (EDC) and Nhydroxysuccinimide (NHS).
Figure 3.5 shows the schematic of the nanoparticle conjugations. After conjugation, the two
functionalized 1pDNA-AuNP-bDNA-NTs for pagA gene and Iel gene were mixed in 1:1 ratio.
The two 2pDNA-coated MNPs were mixed in 1:1 ratio also, for the following multiple detection.
The nanoparticles then were ready for the bio-barcode assay.

3.6 Confirmation of DNA Hybridization
A solution containing tDNA in a PCR tube was put in the thermocycler at 95℃ for 10 min to
separate the dsDNA into ssDNA. Serially diluted DNA samples (40 µL) were mixed with 0.8 mg
MNPs in 200 µL assay buffer (10mM PBS buffer, 0.15M NaCl, 0.1%SDS, pH=7.4). The
hybridization reaction was maintained at a temperature of 45℃ for 45 min in an incubator. After
hybridization, the MNPs with DNA target were washed twice with the assay buffer, then
resuspended in 200 µL of assay buffer. The functionalized AuNPs (100 µL) were centrifuged at
12,000 g for 20 min and the unreacted thiolated oligonucleotides in the supernatant were

44

removed. Finally the purified AuNPs complex was resuspended in 500 µL of assay buffer. The
AuNPs complex (40 µL) was then added into 200 µL solution containing MNPs with DNA
target. The hybridization was incubated at 45 ℃ for 2 h with shaking.
After the sandwich structure (MNP- 2pDNA / Target DNA / 1pDNA-AuNP-bDNA) was
formed, the assay was put on the magnetic separator for 3 min and then the supernatant was
removed. Unreacted solution components (DNA and AuNPs) were washed away 5 times with
500 µL of assay buffer in order to effectively remove AuNPs that were not specifically bound to
the MNPs through hybridization.
Fluorescent detection of bio-barcode assay
A

tDNA

MNPs
2pDNA

with

The composite of
MNP-2pDNA/
tDNA

B

The composite of
MNP-2pDNA/
tDNA/1pDNAAuNP-bDNAfluorescein
bDNA
release

Probe
separation
bDNA-fluoresceins

Magnetic field

Figure 3.6. Schematic of the fluorescent bio-barcode assay: A) formation of MNP2pDNA/tDNA/1pDNA-AuNP-bDNA-fluorescein; B) bDNA-fluorescein separation and release.

45

The MNP-target-AuNP complex was resuspended in 200 µL of 0.5M DTT solution. To
release the bDNA from the surface of AuNPs, the complex was incubated at 50℃ for 15 min,
and then 45 min at 25℃ under vortex. After magnetic separation and centrifugation, the released
bDNA was ready for fluorescent measurement. Figure 3.6 shows a schematic of the fluorescent
bio-barcode assay.
Gold Nanoparticle-based Electrochemical Biosensor Detection
The MNP-target-AuNP sandwich complexes were measured by differential pulse
voltammetry (DPV) on SPCE under the optimum accumulation time. The potential was scanned
from 1.25V to 0.0V with a step potential of 10 mV, modulation amplitude of 50 mV, and scan
rate of 33.5 mV/s (Pumera, Aldavert et al. 2005). Figure 3.7 shows the schematic of the
electrochemical detection of AuNPs. Following oxidative gold metal dissolution in an acidic
solution, the released Au

3+

ions were reduced on SPCE and indirectly quantified by differential

pulse voltammetry (DPV). A potentiostat/galvanostat
Au

3+

3+

Au
3+
Au Au3+
Au
AuNPs
dissolution

DPV

Screen-printed carbon
Electrode (SPCE)
Figure 3.7. Schematics of electrochemical measurement of AuNPs. It is composed of 2 steps:
3+
AuNP dissolution on the SPCE and DPV measurement of Au .

46

with PowerSuite software (Princeton Applied Research, TN) was used for electrochemical
measurement and data analysis.

3.7 Electrochemical Validation and Optimization of the Biosensor
Cyclic voltammetry was performed using a potentiostat/galvanostat (263A, Princeton
Applied Research, MA) in the presence of dissolved oxygen. The screen-printed electrode was
connected to the potentiostat/galvanostat with a specially adapted electrical edge connector. The
scanning potential was between -0.4 V and 0.6 V at a scan rate of 50 mV/s.
Stripping voltammetric measurements were performed with an in situ deposition of the
bismuth film and target metals in the presence of dissolved oxygen. Studies were carried out by
dropping a 100 µL of sample solution in the well. Each electrochemical measurement was
carried out in triplicate. A deposition potential of -1.2 V vs. Ag/AgCl was applied to the carbon
working electrode without solution stirring. After deposition, the voltammogram was recorded
by applying a positive-going square-wave voltammetric potential scan (with a frequency of 20
Hz, amplitude of 25 mV, and potential step of 5 mV). The scan was from -1.2 V to 0.0 V. To
evaluate the reusability of electrochemical sensors, a conditioning step at +0.3 V without
solution stirring was used to remove the target metals and bismuth prior to the next cycle. Bare
SPCE sensors without in situ bismuth film coating were employed for comparison, with
measurement procedures similar to those employed with the bismuth film-coated electrodes
(with the exception of using bismuth (III)). All experiments were carried out at room temperature.

3.8 Electrochemical analysis
Stripping voltammetric measurements were performed with an in situ deposition of the

47

bismuth film and NT ions in the presence of dissolved oxygen. Studies were carried out by
dropping a 100 µL of the sample solution in the well. A deposition potential of -1.2 V vs.
Ag/AgCl was applied to the carbon working electrode without stirring the solution. After
deposition, the voltammogram was recorded by applying a positive-going square-wave
voltammetric potential scan (with a frequency of 20 Hz, amplitude of 25 mV, and potential step
of 5 mV). The scan was from -1.2 V to 0.0 V (Wang, Lu et al. 2000). Based on our previous
optimization studies, 1 mg/L of bismuth and 10 min of deposition time were used for all
SWASV measurements. All experiments were carried out at room temperature. Figure 3.8 shows
the electrochemical test systems.

Personal
computer with
PowerSuite
program

Potentiostat /galvanostat
263A

SPCE Chip

Figure 3.8. The composition of the electrochemical detection system: personal computer with
PowerSuite program, potentiostat/galvanostat 263A, and a SPCE chip.

48

CHAPTER 4: RESULTS AND DISCUSSION

4.1 Characterization of Nanoparticles
The AuNP dimension and spectroscopic properties were characterized by using a
transmission electron microscope (TEM) and a UV-Vis-NIR scanning spectrophotometer
respectively. Figure 4.1A shows a transmission electron microscopy (TEM) image of our
synthesized AuNPs with an average diameter of 15 nm. The dimension of AuNPs is homogenous.
Figure 4.1B shows the absorption peak of the AuNPs at 519 nm wavelength. After one month
storage in the room temperature, the AuNPs did not aggregate and their spectroscopic
absorbance property was stable.

A

50 nm

49

Figure 4.1 (cont’d)

Absorbance (AU)

2.5

B

2
1.5
1
0.5
0
400

450

500

550

600

650

700

750

800

Wavelength (nm)

Figure 4.1. Characterization of AuNPs: A) TEM image of AuNPs, showing an average of 15 nm
in diameter; B) Absorbance spectrum of AuNPs at 519 nm.
Figure 4.2 shows a TEM image of our synthesized CdS nanoparticles with an average
diameter of 7 nm, and PbS nanoparticles with an average diameter of 3 nm. After one month of
storage at 4°C, the CdS and PbS NTs were stable and did not aggregate.

A

B

50 nm

50 nm

Figure 4.2. TEM images of NTs: A) CdS, showing an average diameter of 7 nm; B) PbS,
showing an average diameter of 3 nm.

50

4.2 Functionalization of Nanoparticles
The 6-FAM labeled 2pDNA was used for confirming the functionalization of MNPs. Two
controls were taken for evaluation of the conjugation efficiency. One was only the 6-FAM
pDNA, and the other was 6-FAM pDNA and MNPs without sulfo-SMCC cross-linker. If the
conjugation is efficient, less unconjugated 6-FAM labeled pDNAs will be left in the supernatant
solution after magnetic separation.
Table 4.1 shows that the 6-FAM labeled bDNA has the highest fluorescence signal because
there are no MNPs in the solution. Magnetic separation has no effect on it. Without sulfo-SMCC
as a cross-linker in solution, the pDNA is not conjugated to the MNPs, hence the magnetic
separation will not remove the 6-FAM labeled pDNA. However, some of the pDNA in solution
could be washed out during magnetic separation; hence a decrease in fluorescence signal is
observed. After adding sulfo-SMCC into the conjugation reaction, the conjugation efficiency
improved greatly and the fluorescence signal deceased markedly. The result shows sulfo-SMCC
worked as an efficient cross-linker to conjugate the thiolated oligonucleotides and amine-coated
MNPs. Because the synthesized AuNPs have a peak absorbance at 519 nm, in order to avoid its
interference with the excitation wavelength (520 nm) of 6-FAM, another fluorescent dye (TEX
613) was used to confirm the functionalization of AuNPs. The unconjugated TEX 613 labeled

Table 4.1. Fluorescence signal of supernatant solution for evaluation of conjugation efficiency
between MNPs and 2pDNA.
MNPs + SulfoSMCC + 6-FAM
DNA

MNPs + 6-FAM
DNA (Control 2)

6-FAM DNA
(Control 1)

Fluorescence (0.1s)
(Counts)

72735

293003

368146

Standard deviation

248.7

821.9

1942.1

51

Fluorescence

45
40
35
30
25
20
15
10
5
0

1 nmol barcode without AuNPs
1 nmol bDNA DNA without Au-NPs
1 nmol barcode with AuNPs
1 nmol bDNA DNA with Au-NPs

600

620

640

660

680

700

Wavelength (nm)

Figure 4.3. Fluorescence signal of supernatant solution for evaluation of conjugation efficiency
between AuNPs and bDNA.
bDNA was left in the supernatant solution after centrifugation at 12,000 g for20 min. The control
was the TEX 613 labeled bDNA in the same buffer solution without AuNPs.
Figure 4.3 shows that almost half of the thiolated bDNA has been immobilized on the surface
of An-NPs. The bio-barcoded AuNPs were found to be stable in solution in terms of nonaggregation. This phenomenon is probably because the negatively charged bDNA repels each
other. Hence no aggregation occurs even at long storage time. The solution of bio-barcoded
AuNPs retained its deep red color under room temperature for over a month, demonstrating
stability of conjugation. On the contrary, the AuNPs in the solution without pDNA aggregated to
larger particles and precipitated finally under the same buffer condition.
Confirmation of the functionalization of MNPs and AuNPs was accomplished in our
previous publication (Zhang, Carr et al. 2009). 1-Ethyl-3-[3-dimethylaminopropyl] carbodiimide
Hydrochloride (EDC) and N-hydroxysuccinimide (NHS) were used to crosslink the carboxylic
group on NTs and the amine group on bio-barcoded AuNPs.

52

Under white light

A

B

C

Under UV light

A

B

C

Figure 4.4. Confirmation of conjugation of NTs and AuNPs. Three test tubes labeled A, B, C are
under white light and UV light after centrifugation at 12,000 g for 30 min. (Tube A: biobarcoded AuNPs, NTs, EDC, and NHS; tube B: bio-barcoded AuNPs and NTs; tube C: NTs
only).
Figure 4.4 shows 3 test tubes labeled A, B, C after centrifugation at 12,000 g for 30 min.
Tube A contains bio-barcoded AuNPs, NTs, EDC and NHS. Tube B and C are two controls.

53

Tube B has no cross linker while tube C only contains CdS NTs. There is a pellet that shows
yellowish fluorescence under UV light in tube A. After the centrifugation, the conjugated
particles become larger and are easier to be precipitated. Most of the NTs are conjugated with
AuNPs and are precipitated in the bottom and show the fluorescence signal under UV light. In
tube B, there is a red pellet under white light in the bottom, while the solution shows yellowish
fluorescence under UV light. It shows that without cross linkers, the NTs and bio-barcoded
AuNPs can not be conjugated. The AuNPs are larger and can be precipitated at 12,000 g so that
we can see the red pellet under white light, while the NTs are smaller and still suspend in
solution, showing the fluorescence in the solution under UV light. Tube C shows that
centrifugation does not precipitate the NTs. The results show that conjugation of NTs and biobarcoded AuNPs has been successful.

4.3 Fluorescent Detection Using Bio-barcode Assay
Figure 4.5 shows that the fluorescence signal of released bDNA has an exponential growth
relationship with different concentrations of tDNA. At a concentration of 10 µg/mL tDNA, the
fluorescence signal has an average count of 5711. The signal linearly increases with increasing
concentration of tDNA.

54

Fluorescence of barcode DNA
(0.1sec) (Counts)

7000
6000
5000
4000
3000
2000
1000
0
1E-4
control

1E-3
0.001

0.01

0.1

1

10

Concentration (g/mL)
Figure 4.5. The fluorescence signal of released bDNA with different concentrations of tDNA.
Table 4.2. The relationship between tDNA and the fluorescence signal of released bDNA

Sample Concentration (µg/mL)

10

1

0.1

0.01

0.001

0

Fluorescence reading (counts)

5711

1440

777

426

270

117

3 * Standard Deviation (±)

380.88

209.16

18.52

67.26

28.84

12.01

Table 4.2 shows the mean and variance of the signal. The results show that the assay can
-16

detect as low as 1 ng/mL tDNA (or 2.15 × 10

mol).

55

4.4 Electrochemical Validation and Characterization of SPCE Chips
Cyclic voltammetry of ferricyanide on SPCE chips
Cyclic voltammetry (CV) was performed as the initial electroanalytical technique for the
electrochemical characterization of the sensor surface due to its versatility and relative ease in
measurement. A well characterized redox couple (ferrocyanide/ ferricyanide) was used to
explore the electrochemical nature of the electrode surface. Figure 4.6 shows a cyclic
voltammogram obtained on the electrochemical sensor covered by 100 µL unstirred solution of 5
mM ferricyanide. The ratio of the peak anodic current to the peak cathodic current was 0.95. The
peak separation was much larger than the predicted value of the reversible voltammetric
response and increased with scan rate. The results suggest that voltammetric response on the
electrochemical sensor can be classed as irreversible.

1.50E-04

Current (A)

1.00E-04
5.00E-05
0.00E+00
-5.00E-05
-1.00E-04
-1.50E-04
-0.4

-0.2

0

0.2

0.4

0.6

Potential (V vs. Ag/AgCl)
Figure 4.6. Cyclic voltammogram of bare SPCE sensor in a 5mM K3Fe(CN)6 solution with 1M
KCl. Scan rate is 50mV/s.

56

Reusability test
To avoid the interference from the residue of target metals and/or bismuth during the test of
reusability, various cleaning time (10s, 30s, 1 min) was applied after each SWASV measurement.
After 30 sec of cleaning, there was no oxidative signal of lead(II) and bismuth on the stripping
voltammogram. The result showed that 30 sec was enough to oxidize and remove all bismuth
and metal ions from the surface of the electrochemical sensor. Five milligrams per liter of lead(II)
solution was then used to evaluate the reusability of the bismuth film-coated SPCE sensor and
bare SPCE sensor.
Figure 4.7 shows that the bismuth film coating improves peak current height of SWASV of
lead(II), compared to the bare SPCE sensor. When both electrodes were reused, the peak current
of lead(II) decreased gradually.

100

With bismuth
Without bismuth

Peak current (uA)

90
80
70
60
50
40
30
20
10
0
0

1

2

Times

3

4

5

Figure 4.7. The repeatability test of 1 mg/L of bismuth film-coated SPCE sensor and bare SPCE
sensor. Deposition: -1.2 V for 10 min; SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz,
amplitude 25 mV, and potential step 5 mV, at room temperature and in nonstirred solution.

57

When they were used for 5 times, the signal from bismuth film-coated SPCE sensor was only
66.7% compared to the first measurement; the signal from the bare SPCE sensor was 45.7% of
that at the first time. The results show that the electrochemical sensor, with or without bismuth
film coating, is only good for one-time disposable use in SWASV measurement of heavy metal.
Optimization of deposition time
The following parameters were optimized in order to perform the simultaneous determination
of lead(II) and cadmium(II) on electrochemical sensors: deposition potential, concentration of
bismuth(III) co-deposited in situ with the target metals (lead and cadmium), and deposition time.
Deposition potential was chosen as -1.2V because further reduction of the deposition potential
(to -1.4 V) would result in diminished stripping peak heights, multiple stripping peaks, and
increased background current due to the hydrogen evolution (Reeder and Heineman 1998).
Therefore, deposition potential of -1.2V was used in all subsequent SWASV measurement.
For the co-deposition of bismuth (III) ions with the target metals, the electrochemical sensor
worked as a substrate on which a metallic film of bismuth was formed. The deposition of the
bismuth film depends on its ion concentration and deposition (preconcentration) time. Various
concentrations of bismuth (III) were chosen for co-deposition onto the bare SPCE sensor with 5
mg/L of lead(II) at -1.2 V for 2 min of deposition time. Figure 4.8 shows the lead(II) reoxidation
peak on electrochemical sensors under various concentrations of bismuth ion. The peak currents
increased when the concentrations of bismuth ion increased from 0.1 mg/L to 1 mg/L, because
more bismuth was available for co-deposition with lead(II) and improved the stripping signal of
lead(II). When the concentration of bismuth was increased to 10 mg/L, beyond the

58

0.000035
0.00003
10 mg/L
Current (A)

0.000025

1 mg/L
0.1 mg/L

0.00002
0.000015
0.00001
0.000005
0
-1

-0.9

-0.8

-0.7

-0.6

-0.5

Potential (V vs. Ag/AgCl)

Figure 4.8. Stripping voltammogram of 5 mg/L of lead(II) with various concentrations of
bismuth on electrochemical sensor after 2 min deposition at -1.2 V. SWASV scan from -1.2 V
to 0.0 V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV, at room temperature and
in nonstirred solution.
solubility of bismuth in 0.1 M acetate buffer (pH 4.5), precipitation interfered with the codeposition and lowered the stripping signal of lead(II). So the 1 mg/L of bismuth was used for
subsequent SWASV measurements.
The longer the deposition step, the larger is the amount of analyte available on the electrode
during the stripping analysis. Increasing the deposition time resulted in increased signal (height
of peak current) for both the lead(II) and bismuth stripping waves due to the increased amount of
lead ion and bismuth ion reduced on the electrode surface. Kadara and Tothill reported that the
diffusion of ions was accelerated under stirring condition and deposition times between 60 and
90 s were sufficient to obtain well defined stripping peaks for lead(II) at this solution
concentration (Kadara and Tothill 2004). However, in order to detect a 100 µL sample on an

59

electrochemical sensor, stirring the solution is not practical so that extra deposition time is
needed.
Figure 4.9 shows that the peak currents of lead(II) increased rapidly with deposition time in
the interval from 1 to 45 min. The signal from 10 min deposition is roughly twice compared to
the 1 min deposition. For longer deposition time above10 min, the peak current response of all
electrodes decreased gradually and the wave of lead(II) reoxidation became wider. This result is
potentially due to a change in reference electrode potential and the formation of intermetallic
compounds after excessive deposition. So 10 min of deposition time was used in the
subsequence experiments.

0.00009
0.00008

Current (A)

0.00007
0.00006
1 min
2 min
5 min
10 min
20 min
30 min
45 min

0.00005
0.00004
0.00003
0.00002
0.00001
0
-1.2

-1

-0.8

-0.6

-0.4

Potential (V vs. AgCl)

Figure 4.9. Stripping voltammogram on bismuth film-coated SPCE sensors with various
deposition times. Deposition at-1.2 V; SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz,
amplitude 25 mV, and potential step 5 mV, at room temperature and in nonstirred solution.

60

SWASV measurement of lead and cadmium
Because the unimpeded diffusion to the bismuth-solution interface is available during the
reoxidation step, theoretically, sharper and better defined stripping peaks are expected on
bismuth film-coated SPCE sensors compared to bare SPCE sensors. However, both bismuth
film-coated SPCE sensors and bare SPCE sensors generated sharp and well-defined stripping
peaks in this study. Figure 4.10 shows well-defined sharp peaks over a flat baseline on bismuth
film-coated SPCE sensors, following 10 min of deposition. Peak currents of various lead(II)
concentrations (0.5 mg/L, 2 mg/L and 5 mg/L) are 36 µA, 51 µA, and 75µA, respectively. Peak
currents of various cadmium(II) concentrations (0.5 mg/L, 2 mg/L and 5 mg/L) are 37 µA, 48
µA, and 74 µA, respectively. Bismuth shows a peak current around 39 µA at -0.26 V vs.
Ag/AgCl.
Pb

Cd

0.00008
0.00007

a

Current (A)

0.00006

Bi

0.00005

b

0.00004
0.00003

c

0.00002
0.00001
0
-1.2

-1

-0.8

-0.6

-0.4

-0.2

0

Potential (V vs. Ag/AgCl)

Figure 4.10. Stripping voltammogram of a mixture of lead(II) and cadmium(II) on bismuth filmcoated SPCE sensors (a: 5 mg/L lead(II) and 5 mg/L cadmium(II); b: 2 mg/L lead(II) and 2 mg/L
cadmium(II); c: 0.5 mg/L lead(II) and 0.5 mg/L cadmium(II)). Deposition: -1.2 V for 10 min;
SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV,
with 1mg/mL bismuth, at room temperature and in nonstirred solution.

61

Figure 4.11 shows the stripping voltammograms for lead(II) and cadmium(II) obtained on
bare SPCE sensors. Peak currents of various lead(II) concentrations (0.5 mg/L, 2 mg/L and 5
mg/L) were 20 µA, 40 µA, and 67 µA, respectively. Peak currents of various cadmium(II)
concentrations (0.5 mg/L, 2 mg/L and 5 mg/L) were 15 µA, 30 µA, and 40 µA, respectively.
When the concentration of lead(II) and cadmium(II) was 0.1 mg/L, there was no current peak in
stripping voltammogram on both bare SPCE sensors and bismuth film-coated SPCE sensors
(data not shown). The background noise was around 7 µA for both metal ions on bismuth filmcoated and bare SPCE sensors.
Figure 4.12 shows that the stripping performance for the same concentration of lead(II) on
bare SPCE sensors exhibited a roughly comparable peak to that obtained on bismuth film-coated

0.00008

Pb

0.00007

a

Current (A)

0.00006

Cd

0.00005

b

0.00004
0.00003
0.00002

c

0.00001
0
-1.2

-1

-0.8

-0.6

-0.4

-0.2

0

Potential (V vs. Ag/AgCl)

Figure 4.11. Stripping voltammogram of a mixture of lead(II) and cadmium(II) on bare SPCE
sensors. (a: 5 mg/L lead(II) and 5 mg/L cadmium(II); b: 2 mg/L lead(II) and 2 mg/L cadmium(II);
c: 0.5 mg/L lead(II) and 0.5 mg/L cadmium(II)). Deposition: -1.2 V for 10 min; SWASV scan
from -1.2 V to 0.0 V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV, with 1mg/mL
bismuth, at room temperature and in nonstirred solution.

62

100

Lead(II) on bismuth film-coated SPCE
Lead(II) on bare SPCE
Cadmium(II) on bismuth film-coated SPCE
Cadmium(II) on bare SPCE

90
Peak current (uA)

80
70
60
50
40
30
20
10
0
0

1

2
3
4
Concentration of metal ions (mg/L)

5

Figure 4.12. The peak currents of stripping voltammogram with different concentrations of
lead(II) and cadmium(II).
SPCE sensors. However, the stripping performance for the same concentration of cadmium(II)
on bare SPCE sensors exhibited a much lower peak to that of bismuth film-coated SPCE sensors.
When the signal/noise was greater than 3, we concluded that the detection limits of cadmium(II)
were 0.5 mg/L on bismuth film-coated SPCE sensors and 2 mg/L on bare SPCE sensors in 100
µL of sample solution. The detection limits of lead(II) were 0.5 mg/L on bismuth film-coated
SPCE sensors and 0.5 mg/L on bare SPCE sensors in 100 µL of sample solution. Cadmium(II)
detection benefited from the addition of bismuth.

4.5 Detection of AuNPs on SPCE Chips
CV of unmodified AuNPs on SPCE

63

Compared with conventional electrodes, screen-printed carbon electrodes have several
advantages, such as simplicity, convenience, low cost and the avoidance of contamination
between samples (Susmel, Guilbault et al. 2003; Miscoria, Desbrieres et al. 2006). Biosensors
based on screen-printed carbon electrode (SPCE) have been extensively used for the detection of
glucose (Guan, Li et al. 2005; Xu, Li et al. 2005), cholesterol (Carrara, Shumyantseva et al.
2008), antigens (Kim, Seo et al. 2006) and DNA (Ruffien, Dequaire et al. 2003; HernandezSantos, Diaz-Gonzalez et al. 2004).
Oligonucleotide-functionalized AuNPs were used as signal indicators in this study because of
their ease of fabrication, greater oligonucleotide binding capabilities, stability under a variety of
0.00015

A

0.0001

B

Current (A)

0.00005

0
0
-0.00005

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

Potential (V vs. Ag/AgCl)

-0.0001

-0.00015

-0.0002

-8

Figure 4.13. Cyclic voltammogram of unmodified AuNPs (1.17 x 10 M) on SPCE. : (A)
AuNPs in 0.1M HCl; (B) 0.1 M HCl. CV scan from 1.4V to 0.0, scan rate 100 mV/s after 1.25 V
electrooxidation for 2 min.

64

conditions, and good electrochemical properties (Demers, Ostblom et al. 2002; Lytton-Jean and
Mirkin 2005).
Figure 4.13B shows that when there is no AuNPs, the carbon electrode is oxidized when
performing oxidative procedure at 1.25V for 2 min. The carbon electrode can produce a
reduction peak at 0.58 V. When the AuNPs exist in the solution, the AuNPs are oxidized instead
of the carbon electrode under 1.25V for 2 min. The curve (Figure 4.13A) shows that there is a
3+

reduction peak of Au

at 0.35 V. Meanwhile, the background signal at 0.58 V from the carbon

electrode is suppressed greatly by AuNPs. The possible reason is that the surface of SPCE is
covered by a self-assembled layer of AuNPs due to physical adsorption. The SPCE is protected
by AuNPs and this prevents the electrooxidation process from proceeding on the carbon
electrode surface.
Optimization of detection of probe modified-AuNPs
The accumulation time is the time from dropping the sample on the SPCE to the beginning of
electrochemical detection. It is an important factor for the electrochemical measurement of
AuNPs because more AuNPs would be adsorbed on the SPCE surface when the accumulation
time is longer. The electrooxidation process would generate more Au

3+

ions in the solution,

which could be translated to a higher DPV signal and thus improve the biosensor sensitivity.
The sample solution was dried after 20 min at room temperature. The drying allowed most of
the AuNPs to adsorb physically on the SPCE surface. Then 100 µL of 0.1M HCl was added
again for DPV measurement. Figure 4.14 shows the DPV current for 0, 10, 20 min of
accumulation times. Figure 4.14A has 20 min of accumulation time which allows for more
AuNPs to be reduced or oxidized, leading to a comparatively high signal. With 10 min and 0 min
accumulation times, peak current are at 0.86 mA and 0.36 mA, respectively. They are only 72%

65

0.00015

A
0.0001

B
C

Current (A)

0.00005
0

D

-0.00005
-0.0001
-0.00015
-0.0002
0

0.5
1
Potential (V vs. Ag/AgCl)

1.5

Figure 4.14. Cyclic voltammogram of probe-modified AuNPs in 0.1 M HCl on SPCE: (A)
accumulation for 20 min; (B) accumulation for 10 min; (C) no accumulation; (D) control, no
AuNPs and no accumulation. Conditions: electrooxidation potential, +1.25 V; electrooxidation
time, 2 min, CV scan from +1.4 V to 0 V, scan rate 100 mV/s, nonstirred solution.
and 30% of the reduction peak current at 1.21 mA for the 20 min accumulation time. Drying led
to more AuNPs available on the surface of SPCEs, resulting in the higher sensitivity of SPCE
biosensors. Accumulation time of 20 min was used for the succeeding DPV detection of the
DNA sandwich complex.
DPV detection of DNA sandwich complex
Figure

4.15

shows

the

DPV

response

of

the

sandwich

complex

(MNP-

2pDNA/tDNA/1pDNA-AuNPs) after hybridization for various tDNA concentrations (0.7-700

66

0.00004

b

Current (A)

0.000035
0.00003

Peak current at 0.4V (A)

a

0.000045

0.00005
0.00004
0.00003
0.00002
0.00001
0
0.0001

c

0.000025

y = 5E-06Ln(x) + 4E-05
R2 = 0.9937

0.001

0.01

0.1

1

Target DNA concentration (µg/m L)

0.00002
0.000015

d

0.00001
0.000005
0
0

0.2

0.4

0.6

0.8

1

1.2

1.4

Potential (V vs. Ag/AgCl)
Figure 4.15. DPV hybridization response of different concentrations of DNA targets on SPCE (a:
700 ng/mL; b: 70 ng/mL; c: 7 ng/mL; d: 0.7 ng/mL) and the calibration plot between peak
current and the tDNA concentration (inset). DPV scan from +1.25 V to 0 V, step potential 10
mV, modulation amplitude 50 mV, scan rate 50 mV/s, and nonstirred solution.
3+

ng/mL). Following oxidative gold metal dissolution in the acidic solution, the Au

was reduced

at the potential around 0.4V (vs. Ag/AgCl) on the SPCE. Inset shows that the peak current has a
log-linear relationship with increasing concentrations of tDNA. The detection limit is 0.7 ng/mL
of tDNA.

4.6 Biosensor Detection of Single Target DNA

67

Figure 4.16 is a schematic showing the bio-barcode assay. After immobilization of
oligonucleotides on the surface of nanoparticles, both nanoparticles can bind with the tDNA to
form a sandwich structure, due to the specificity of pDNA. Figure 4.16A shows the tDNA
sample hybridizing with the pDNA on the MNPs, forming a MNP-2pDNA/tDNA complex
(middle picture). Then bio-barcoded AuNPs are added to form a sandwich structure consisting of
MNP-2pDNA/tDNA/1pDNA-AuNP-bDNA (picture on the right). After hybridization is
complete, the NTs on the bDNA are released in 1M nitric acid and the metal ions are measured
with SWAPV on the SPCE DNA sensor (Figure 4.16B).

A

tDNA

MNPs with
2pDNA

The composite of
MNP-2pDNA/
tDNA/1pDNAAuNP-bDNA-NTs

The composite of
MNP-2pDNA/
tDNA

B

bDNA
release

tDNA
separation

M

2+

2+

M

2+ 2+
NTs
M M
dissolut
2+ 2+
M M
ion
bDNA-NTs

Magnetic field

Figure 4.16.
Schematic of the bio-barcode assay. (A) formation of MNP-2pDNA
/tDNA/1pDNA-AuNP-bDNA-NTs sandwich structure; (B) tDNA separation and tracer
dissolution.
68

0.0001

2 ng/mL

0.00009
0.00008

0. 2 ng/mL

0.00007

20 pg/mL
Control

0.00006
0.00005

2 pg/mL

0.00004

0.2 pg/mL
20 fg/mL
2 fg/mL

Current (A)

A

0.00003
0.00002
0.00001

-1.2

-1

-0.8

-0.6

0
-0.2

-0.4

Potential (V vs. Ag/AgCl)

120

B

Current (uA)

100
80
60
40
20
0
1E-06 0.00001 0.0001

0.001

0.01

0.1

1

10

Target DNA Concentration (ng/mL)
Figure 4.17. Single detection of various concentrations of pagA gene from Bacillus anthracis
with the bio-barcode biosensor using PbS NTs. (A) stripping voltammogram of
various
concentrations of pagA gene from Bacillus anthracis; (B) the relationship of tDNA concentration
and peak current. Conditions: deposition at -1.2 V for 10 min; SWASV scan from -1.2 V to 0.0
V, frequency 20 Hz, amplitude 25 mV, and potential step 5 mV, with 1mg/mL bismuth, at room
temperature and in nonstirred solution.

69

2+

Figure 4.17 shows the electrochemical signal of Pb

from pagA gene from Bacillus

anthracis in single target detection. The peak currents of various tDNA concentrations (2 ng/mL,
0.2 ng/mL, 20 pg/mL, 2 pg/mL, 0.2 pg/mL, 0.02 pg/mL, 2 fg/mL, and 0.2 fg/mL) are 96 µA, 76
µA, 63 µA, 46 µA, 21 µA, 14 µA, and 13 µA respectively. The biosensor shows that the
sensitivity of single detection of pagA gene from Bacillus anthracis is 0.2 pg/mL or 200 fg/mL.

4.7 Biosensor Detection of Multiplex Target DNA
Figure 4.18A is a schematic showing the multiple bio-barcode assay. After immobilization of
oligonucleotides on the surface of nanoparticles, both nanoparticles can bind with the tDNAs to
form a sandwich structure due to the specificity of pDNA. First, the tDNA samples hybridize
with the pDNA on the MNPs, forming a MNP-2pDNA/tDNA complex (middle picture). Then
the 1pDNA-AuNP-bDNA-NTs complex is added to form a sandwich structure consisting of
MNP-2pDNA/ tDNA/1pDNA-AuNP-bDNA-NTs. After hybridization is complete, the NTs on
the bDNA are released in 1M nitric acid and the NT ions are measured by SWASV on the SPCE
chip. Figure 4.18B shows that SWASV measurements are performed with an in situ deposition
of the bismuth film and target metals (Pb and Cd) in the presence of dissolved oxygen using a
potentiostat/galvanostat.

70

Biobarcoded
AuNPs
tDNA

MNPs2pDNA

The composite of
MNP-2pDNA/tDNA
Washing

Electrochemical 2+ 2+
2+
Pb Cd Pb
detection
2+ 2+
2+
Cd Pb Cd

The composite of
MNP-2pDNA/
tDNA/1pDNAAuNP-bDNANTs

NTs
dissolution

Magnetic field

Readout

(A)
NT
dissolution

Pb

2+

Pb

2+

2+

Bi

Screen-printed
carbon Electrode
(SPCE)

Deposition

Cd

2+

Cd

2+

Pb

2+

Cd

3+

Cd

Pb

Pb

Bi

Bismuth-coated
SPCE

2+
2+

Cd

2

2+

Pb Cd
Pb

2+

2+

Cd

Anodic
stripping

(B)
Figure 4.18. Schematic of the multiple bio-barcode biosensor: A) bio-barcode assay; B) SWASV
measurement.

71

Figure 4.19 shows the electrochemical signal of released NT ions (Pb

2+

2+

and Cd ) and the

specificity of the biosensor. When there is only 0.05 μg/mL of pagA gene from Bacillus
anthracis, the stripping voltammogram shows a current peak of 39 µA at -0.61 V from dissolved
2+

Pb , and no peak at -0.87 V (Figure 4.19A). When there is only 0.05 μg/mL of the insertion
element (Iel) gene of Salmonella Enteritidis, the stripping voltammogram shows a current peak
2+

of 46 µA at -0.87 V from dissolved Cd , and a small peak at -0.61 V (Figure 4.19B).
A
Iel gene

0.00004
0.00003
0.00002
0.00001

control

0
-1.2

-0.7

B
pagA gene

0.00006
Current (A)

Current (A)

0.00005

0.00005
0.00004
0.00003
0.00002
0.00001

control

0
-1.2

-0.2

-0.7

-0.2

Potential (V vs. Ag/AgCl)

Potential (V vs. Ag/AgCl)
0.00006

C

Current (A)

0.00005

Iel gene

0.00004

pagA gene

0.00003
0.00002
0.00001
0
-1.2

control
-1

-0.8

-0.6

-0.4

-0.2

0

Potential (V vs. Ag/AgCl)

Figure 4.19. Specificity test of the multiple bio-barcode biosensor using distilled water as control.
(A) control and 0.05 μg/mL of the insertion element (Iel) gene of Salmonella Enteritidis; (B)
control and 0.05 μg/mL of pagA gene from Bacillus anthracis; (C) control and a mixture of 0.05
μg/mL of the insertion element (Iel) gene and 0.05 μg/mL of pagA gene. Conditions: deposition
at -1.2 V for 10 min; SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz, amplitude 25 mV,
and potential step 5 mV, with 1mg/mL bismuth, at room temperature and in nonstirred solution.

72

The possible reason would be that the washing steps were not thorough and the unreacted
PbS NTs generate the noise. When the sample contains no tDNA, there are no stripping signal at
2+

2+

-0.61 V (Pb ) and -0.87 V (Cd ). When the sample contains 0.05 μg/mL of the pagA gene
from Bacillus anthracis and the insertion element (Iel) gene of Salmonella Enteritidis, the peak
current at -0.61 V (vs. Ag/AgCl) is 35 µA and the peak current at -0.87 V (vs. Ag/AgCl) is 39
µA, respectively (Figure 4.19C). The results show that this bio-barcoded DNA sensor has a good
specificity. It can differentiate pagA gene from Bacillus anthracis (labeled with CdS) and Iel
gene of Salmonella Enteritidis (labeled with PbS) when the concentration of tDNA is relatively
high.
Figure 4.20 shows the sensitivity of the biosensor. For the detection of the pagA gene from
Bacillus anthracis, the peak currents at -0.61 V (vs. Ag/AgCl) of various concentrations (50
ng/mL, 5 ng/mL, 500 pg/mL, and 50 pg/mL) are 38 µA, 34 µA, 29.5 µA, and 23µA, respectively.
For the detection of the insertion element (Iel) gene of Salmonella Enteritidis, the currents at 0.87 V at various tDNA concentrations (50 ng/mL, 5 ng/mL, 500 pg/mL, and 50 pg/mL) are 42.5
µA and 35 µA, 32 µA, and 27 µA, respectively. Figure 4.20B shows that the stripping signal of
released Pb

2+

and Cd

2+

have a linear relationship with the logarithmic concentrations of tDNA.

The signal increases with increasing logarithmical concentration of tDNA. The results show that
the detection limit of this bio-barcoded DNA sensor is as low as 50 pg/mL (lowest concentration
tested) of tDNAs using CdS and PbS NTs.

73

A

0.00006
0.00005

50 ng/mL
0.5 ng/mL

Current (A)

5 ng/mL

0.00004
0.00003
0.00002

50 pg/mL

Control

0.00001
0
-1.2

-1

-0.8

-0.6

-0.4

-0.2

0

Potential (V vs. Ag/AgCl)

B

50
45
40

y = 2.1498Ln(x) + 47.99
R2 = 0.9709

35
Current (uA)

30

y = 2.1498Ln(x) + 44.99
R2 = 0.9865

25
20

Bacillus anthracis

15

Salmonella Enteritidis

10
5
0

0.00001

0.0001

0.001

0.01

0.1

1

Target DNA Concentration (ug/mL)

2+

2+

Figure 4.20. The stripping signal of released NT ions (Pb and Cd ) with different
concentrations of tDNAs: A) stripping voltammogram of various concentrations of tDNA;. B)
the calibration plot between peak current and the tDNA concentration. Conditions: deposition at
-1.2 V for 10 min; SWASV scan from -1.2 V to 0.0 V, frequency 20 Hz, amplitude 25 mV, and
potential step 5 mV, with 1mg/mL bismuth, at room temperature and in nonstirred solution

74

4.8 Conclusion and Summary
Conclusion
A highly amplified, nanoparticle-based, bio-barcoded electrochemical biosensor for the
multiple detection of pagA gene (accession number = M22589) from Bacillus anthracis and
insertion element (Iel) gene (accession number = Z83734) from Salmonella Enteritidis was
described in this research. The biosensor system was mainly composed of three nanoparticles:
gold nanoparticles (AuNPs) for a carrier of signals, magnetic nanoparticles (MNPs) for clean
separation and preconcentration, and nanoparticle tracers (NTs, such as PbS and CdS) for signal
amplification. After mixing the nanoparticles with the tDNA, the sandwich structure (MNP2pDNA / tDNA / 1pDNA-AuNP-bDNA-NTs) was formed. The released NT ions showed a nonoverlapping stripping behavior by square wave anodic stripping voltammetry (SWASV) on
screen-printed carbon electrode (SPCE) chips. The screen-printed carbon electrode (SPCE) was
chosen because of its broad potential window, low background current, rich surface chemistry,
low cost, chemical inertness, and suitability for the detection of trace levels of metal ions.
SWASV has an extremely favorable signal-to-background ratio and remarkable sensitivity due to
an effective pre-concentration step of anodic stripping techniques. Bio-barcode assay has been
shown to have extraordinarily PCR-like sensitivity because the large ratio between thiolated
single-strand oligonucleotide barcodes and pDNA on AuNPs provides significant amplification.
There were six specific aims of this study as follows:
1) Synthesize and characterize gold nanoparticles and nanoparticle tracers (PbS and CdS):
The nanoparticles were synthesized successfully and characterized. The absorption peak of the
AuNPs was at 519 nm wavelength. TEM shown the diameters of AuNPs, PbS NTs and CdS NTs
were 15 nm, 3 nm, and 7 nm respectively. They were stable in room temperature for months.

75

2) Functionalize gold nanoparticles and magnetic nanoparticles, and conjugate bio-barcoded
AuNPs with nanoparticle tracers: The conjugation reaction between the two nanoparticles and
thiolated oligonucleotides, carboxylic group on NTs and amine group on bio-barcoded AuNPs
were efficient.
3) Evaluate and optimize SPCE chips for NT ions detection: The cyclic voltammogram of
ferricyanide showed that the voltammetric response of SPCE chips was irreversible. The
electrochemical sensors were only good for one-time use. Due to the limitation of stirring,
deposition potential of -1.2 V, 1 mg/L of bismuth and 10 min deposition time were applied to the
all SWASV measurements.
4) Fluorescence detection of bio-barcode assay and gold nanoparticle detection to confirm
hybridization: The fluorescence signal of released bDNA had an exponential relationship with
tDNA concentration. The detection limit of this bio-barcoded DNA assay was 1 ng/mL (or 2.15
-16

× 10

mol). After hybridization, the detection limit of the insertion element (Iel) gene of
-16

Salmonella Enteritidis was 0.7 ng/mL (or 1.5× 10

mol) using AuNPs as an electrochemical

indicator.
5) Evaluate biosensor sensitivity using a single purified PCR product from target organisms:
The sensitivity of single detection of pagA gene from Bacillus anthracis using PbS NTs is as low
as 0.2 pg/mL.
6) Design multiplex detection of multiple targets in the same samples and evaluate its
sensitivity and specificity: The biosensor had a good specificity, and the detection limit of this
multiplex bio-barcoded DNA sensor was 50 pg/mL using PbS or CdS NTs.
Limitation and Future Possibilities

76

The nanoparticle-based bio-barcoded DNA sensor faces several issues that need to be
addressed before it can be adopted as an on-site biosensor. Though the detection system is easy
to miniaturize, the current system is still large and heavy; Even though this detection system can
reach the PCR-like sensitivity, DNA extraction from pathogens and restriction enzymes to cut
the long gene sequence are still necessary. It will need battery-operated centrifuge.
To further improve the sensitivity of the biosensor, it is important to optimize the ratio of
1pDNA to bDNA to maximize the amplification while maintaining the hybridization efficiency
to the tDNA. A better understanding of the steric hindrance of nanoparticle tracers during
hybridization is essential to improve the hybridization efficiency.
In the future, the SPCE chips can be incorporated into a pocket PC. The biosensor detection
systems will be handheld, portable, cost-effective, have PCR-like sensitivity, can simultaneously
detect multiplex pathogens. Figure 4.21 shows a schematic of the future ultrasensitive
electrochemical biosensor.

Pocket PC
Handheld potentiostat/galvanostat

SPCE

Figure 4.21. Schematic of ultra-sensitive portable electrochemical biosensor system.

77

APPENDICES

78

APPENDIX I
DNA Extraction Procedure (Cited from manufacturer’s manual)

1)

In the biosafety cabinet, pipette 1 ml of bacterial culture into a 1.5 ml micro centrifuge tube,
and centrifuge for 5 min at 5000 x g (7500 rpm).

2)

Remove 910 µL supernatant, then add Buffer ATL 90 µL(supplied in the QIAamp DNA
Mini Kit), to a total volume of 180 μL.

3)

Add 20 μL Proteinase K, mix by shaking, and take it out from biosafety cabinet. Incubate
and shake in hybridization oven at 56°C until the tissue is completely lyses. Vortex
occasionally during incubation to disperse the sample.

4)

Briefly centrifuge the 1.5 ml microcentrifuge tube for 1 min to remove drops from the
inside of the lid. Add 200 μL Buffer AL to the sample, mix by pulse-vortexing for 15 s, and
incubate in hybridization oven at 70°C for 10 min.

5)

Briefly centrifuge the 1.5 ml microcentrifuge tube for 1 min to remove drops from inside
the lid. Add 200 μL ethanol (96–100%) to the sample, and mix by pulse-vortexing for 15 s.

6)

After mixing, briefly centrifuge the 1.5 ml microcentrifuge tube to remove drops from
inside the lid. Carefully apply the mixture (including the precipitate) to the QIAamp Spin
Column (in a 2 ml collection tube) without wetting the rim. Close the cap, and centrifuge
at 6000 x g (8000 rpm) for 1 min.

7)

Place the QIAamp Spin Column in a clean 2 ml collection tube (provided), and discard the
tube containing the filtrate.

8)

Carefully open the QIAamp Spin Column and add 500 μL Buffer AW1 without wetting
the rim. Close the cap, and centrifuge at 6000 x g (8000 rpm) for 1 min.

9)

Place the QIAamp Spin Column in a clean 2 ml collection tube (provided), and discard the
collection tube containing the filtrate.

10) Carefully open the QIAamp Spin Column and add 500 μL Buffer AW2 without wetting
the rim. Close the cap and centrifuge at full speed (20,000 x g; 14,000 rpm) for 3 min.
11) Place the QIAamp Spin Column in a new 2 ml collection tube (not provided) and discard
the collection tube containing the filtrate. Centrifuge at 20,000 x g (14,000 rpm) for 1 min.
12) Place the QIAamp Spin Column in a clean 1.5 ml microcentrifuge tube (not provided), and
discard the collection tube containing the filtrate.

79

13) Carefully open the QIAamp Spin Column and add 200 μL Buffer AE or distilled water.
Incubate at room temperature for 5 min, and then centrifuge at 6000 x g (8000 rpm) for 1
min.
14) The filtrate in the 1.5 mL is the extracted DNA sample. Its concentration and purification
can be measured with SmartSpec 3000 Spectrophotomete.
15) It should be stored at -20°C

80

APPENDIX II
PCR Amplification (Cited from manufacturer’s manual)

1) Thaw the PCR Master Mix at room temperature. Vortex the Master Mix and then spin it
briefly in a microcentrifuge to collect the material in the bottom of the tube.
2) Prepare one of the following reaction mixes on ice:
Component
PCR Master Mix, 2X
Reverse primer, 10µM
Forward primer, 10µM
DNA template
Nuclease-Free Water to

Volume
50µl
1.0–10.0µl
1.0–10.0µl
1–5µl
100µl

Final Conc.
1X
0.1–1.0µM
0.1–1.0µM
<250ng
N.A.

3) Put the PCR tube into the thermocycler.
4) Choose the program name “SALMO” and start PCR amplification.
Denaturation :

94℃ 5 min
94℃ 30 sec
Annealing:
59℃ 30 sec
Extension:
72℃ 45 sec
Cycle Number: 30
Final extension: 72℃ 10 min
Refrigeration: 4℃
After PCR, run the electrophoresis gel or store under -20℃freezer.

81

APPENDIX III
Procedure of Bio-Barcode Assay (Adapted from Hill and Mirkin 2006)
1. 13 nm Gold nanoparticles preparation
1) Clean all glassware with pure water, rinse copiously with pure water and dry in oven at
100℃. Wash a large stir bar.
2) Blow out glassware with N2 or air
3) Prepare 50 ml of 1mM hydrogen tetrochloroaurate (III) trihydrate with pure water
(0.0197 g + 50 ml H2O) in a 100 ml flask. Use plastic spatula.
4) Pour the gold solution into the flask and bring to a vigorous boil while stirring. Using a
foil to cover the opening roughly.
5) While the gold solution is heating, prepare 5 ml of 38.8 mM sodium citrate dehydrate
with pure water in the (0.057g sodium citrate + 5 ml H2O)
6) Once the gold solution is refluxing vigorously, add all the sodium citrate solution and
reseal the flask top with foil paper. The solution will turn from yellow to clear, to black,
to purple and finally deep red.
7) After 15 min, turn heat off and allow the reaction to cool to room temperature.
8) The UV–vis absorption spectrum of AuNPs solution showed a strong surface plasma
resonance at a wavelength of 519 nm.
2. Magnetic nanoparticle preparation
1) Melt the solid of 1,6-hexanediamine to liquid phase at 45C in a water bath for 2 hours
2) A solution of 1,6-hexanediamine (6.5 g), anhydrous sodium acetate (2.0 g) and
FeCl3·6H2O (1.0 g) in Ethylene glycol (30 mL)was stirred vigorously at 50 ℃ to give a
transparent solution.
3) This solution was then coved with an aluminum foil and transferred into an oven and
reacted at 198 ℃ for 6 h.
4) Under ultrasonic conditions, the magnetite nanoparticles were then rinsed with ethanol
and water for 15 min to effectively remove the solvent and unbound 1,6-hexanediamine,
repeat the washing step 3 times respectively. During each rinsing step, the nanoparticles
were separated from the supernatant by using magnet for 3 min.

82

5) The magnetite nanoparticles dried at 50 ℃ in the hybridization oven before
characterization and application.
6) Story the MNPs in storage buffer (Na2HPO4 8.0 mM, NaH2PO4 2mM, NaCl 0.15M,
Tween 20 0.05%, pH 7.4) under 4℃.

4. DNA functionalization on gold Nanoparticles (AuNPs)
1) Take out 5 nmol of thiolated barcode DNA and 0.05 nmol 1pDNA from freezer and mix
them together.
2) Prepare 1 ml of 0.1 M DTT (0.0154g) solution in the disulfide cleavage buffer.
3) Add 100 µL of DTT solution to the barcode DNA, wrap in foil and let stand at room
temperature for 2 hours. Vortex once half an hour.
4) Flushing a Nap-5 column with pure water. 3 column volumes of water must flush through
before adding barcode DNA
5) Adding 100 µL of DNA to the column after all the water has run through
6) Once the 100 µL of DNA has flowed into the column, add 400 µL of pure water to the
column and allow it to flow through uncollected.
7) Then add 950 µL of pure water to the column, and collect the flow through in 1.5 mL
microcentrifuge tubes.
8) Use Nanodrop to determine the DNA concentration.
9) Add 1 ml of AuNPs (concentration: AU = 1 at 520 nm) in a 15 ml test tube.
10) Add the freshly reduced thiolated oligo to the AuNPs. Recode the volume.
11) Wrap in foil and place on shaker overnight at room temperature.
12) Add 195 µL of phosphate adjustment buffer. Calculation: (1000µL AuNP + X µL DNA)
/ 10 = Y µL
13) Add 21.5 µL of surfactant solution. Calculation: (1000µL AuNP + X µL DNA + Y µL
phosphate adjustment buffer) /100 = Z µL
14) Rewrap in foil and place on shaker for 30 min.

83

15) Add salting buffer at six times in 2 days. So every time just add 53.5 µL of salting buffer.
Do the additions while shaking gently. Calculation: (1000µL AuNP + X µL DNA + Y µL
phosphate adjustment buffer) * 0.3/ 2 = S µL.
16) After the last salt addition, allow the particles to equilibrate overnight.
The particles can be store at room temperature for one month in this state.

5. DNA functionalization of magnetic nanoparticles (MNPs)
1) Sonicate MNPs stock solution (50mg/mL) for 30 min to disperse Nanoparticles evenly.
2) Take out 5 mg (100 µL) of the polyamine-functionalized MNPs from stock solution
(50mg/mL). It will react with 0.5 mg of sulfosuccinimidyl 4-N-maleimidomethyl
cyclohexane-1-carboxylate (Sulfo-SMCC) bifunctional linker in 10 mL coupling buffer.
3) Put the tubes into shaker for 1 hour at room temperature.
4) Put it on the magnet 3 min, remove the supernatant (unreacted Sulfo-SMCC) and rinsing
it with the coupling buffer. Repeat the process 3 times. Finally take off all supernatant.
5) Take out 10 nmol of thiolated 2pDNA from freezer.
6) Prepare 1 ml of 0.1 M DTT (0.0154g) solution in the disulfide cleavage buffer.
7) Add 100 µL of DTT solution to 2pDNA, wrap in foil and let stand at room temperature
for 2 hours. Vortex once half an hour.
8) Flushing a Nap-5 column with coupling buffer. 3 column volumes of coupling buffer
must flush through before adding barcode DNA
9) Adding all DNA (suppose the volume now is X) to the column after all the coupling
buffer has run through.
10) Once the X µL of DNA has flowed into the column, add 500-X µL of coupling buffer to
the column and allow it to flow through uncollected.
11) Then add 950 µL coupling buffer to the column, and collect the flow through in 1.5 mL
microcentrifuge tubes.
12) Use Nanodrop to determine the DNA concentration.
Attention: Coupling buffer as a blank.
13) Put the cleaved and purified thoilated oligonucleotide with sulfo-SMCC-modified MNPs
in 1 ml coupling buffer for overnight (>8 h).

84

14) Put it on the magnet for 3 min, remove the supernatant and rinsing it with the coupling
buffer. Repeat the process 3 times.
15) The particles were centrifuged (Eppendorf Centrifuge 5415D) (4,000 RPM; 1 min) and
washed (3 x) with passivation buffer (2x) and then with a storage buffer (2 x) (10 mM
sodium phosphate buffer pH 7.4 with 0.20 M NaCl).
16) Finally, the target binding DNA-modified MNPs were re-dispersed in 2 mL of 0.1 M
PBS solution prior to use.
6. Hybridization of sandwich structure
1) Determine the number of samples to be tested, including one for the negative control
(pure water).
2) Wash the MNPs twice with the assay buffer.
3) A solution (40µL per sample test) containing tDNA in PCR tube was put in the
thermocycler at 95C for 10 min to separate the dsDNA to ssDNA, then put in freezer (20C) immediately for 5 min to keep them separate.
4) Mix in 1.5 mL microcentrifuge tubes. 40 µL DNA sample + 0.8 mg MNPs + X µL assay
buffer = 200 µL
5) Shake the reactions at a temperature 45C for 45 min in the rotating hybridization oven.
So that the MNPs do not settle.
6) During the incubation, centrifuge 200 µL of AuNPs at 13000 g for 30 min. remove the
supernatant and resuspend in 1 mL assay buffer. Repeat 3 times.
7) After final spin/wash, resuspend the AuNPs in assay buffer to generate 1 nM solution
(AU=0.27). Calculation based on the Beer’s law A = alc. The molar absorptivity at 519
8
-1
-1
nm is 2.7 * 10 liter mole cm so that 1 AU = 3.7 nM.
8) Wash the MNPs with target twice with the assay buffer on the magnetic separator, and
finally resuspend in 200 µL assay buffer.
9) Add 40 µL of AuNPs probe recently washed.
10) Incubate at 45C for 1.5 h with in the rotating hybridization oven.

7. Releasing barcode DNA with dithiothreitol (DTT)
1) Prepare a 0.0771 g DTT solution in 1 mL assay buffer (0.5M). (Must be fresh every day )

85

2) Wash the MNP-target-AuNP complexes twice with the assay buffer on the magnetic
separator, and finally suspend the MNP-target-AuNP complexes in 200 µL 0.5M DTT
buffer.
3) Incubate samples at 50℃ for 15 min, and then 45 min at 25℃ under vortex every 5 min.

8. Detection of Barcode DNA
A. conventional fluorophore detection methods (Fluorescence-based barcode DNA)
1) After barcode release, extract the MNPs down to the magnet for 3 min and transfer 150
µL supernatant to clean 1.5 mL microcentrifuge tubes.
2) Spin the supernatants for 5 min at 13,000g to pellet the aggregated AuNPs
3) Prepare serial dilution of known concentrations in barcode release buffer to generate a
calibration curve
4) Remove 100 µL supernatant from gold pellet and place in a new micro-centrifuge tube.
5) Fill the microplate with the 100 µL supernatant.
6) Follow the instrument manual, measure florescence and perform data analysis.
B. Electrochemical detection of barcode DNA

86

APPENDIX IV
Table 5. Buffer preparation.

Buffer
Coupling buffer
Disulfide cleavage buffer
Storage buffer
Passivation buffer
Salting buffer
Phosphate adjustment
buffer
Assay buffer
Surfactant solution

Na2HPO4
(g)
4.392
11.468
0.570
10.119
0.0562

NaH2PO4
(g)
2.291
0.509
0.118
0.449
0.0125

5.884
4.383
5.844

0.562
0.562

0.125
0.125

4.383

NaCl
(g)
5.884

Na2HPO4

Disodium hydrogen phosphate

NaH2PO4
NaCl
SDS

Sodium dihydrogen phosphate
Sodium chloride
Sodium dodecyl sulfate

87

SDS
(g)

0.500
10.000

Pure water
(ml)
500
500
500
500
50
50
500
90

pH
7.2
8.0
7.4
8.0
7.0
7.0
7.4

APPENDIX V
Conjugation of carboxylic group on NTs with amine group on bio-barcoded AuNPs
1. Weigh out 5mg EDC into tube
2. Immediately add 25 µL of CdS-COOH or PbS-COOH to EDC
3. Immediately add 50µL of NHS-DMSO solution (44mg of NHS in 500 µL DMSO)
4. Add 775 µL of pH5 water to make total volume 850µL
5. Quick vortex, quick spin
6. Wrap in tin foil, react 20min at room temp
7. Adjust pH to 7.0 with NaCO3. (4µL of 1M NaCO3 pH8)
8. Add AuNP-barcode-NH2 (61 µL)
9. Wrap in tin foil, tape to vortex at 800rpm for 1hr
10. Add 3µL of 1M NaCO3 pH8
11. Wrap in tin foil, tape to vortex at 800rpm overnight (or 1h)
12. Spin 15min at 12,000 g, wash 3x times with PBS buffer
13. Resuspend in 1000µL of PBS buffer

88

BIBLIOGRAPHY

89

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