Monitoring DNA hybridization using
Rasheeda M. Hawk,1Maria V. Chistiakova,1and Andrea M. Armani1,2,*
1Mork Family Department of Chemical Engineering and Materials Science, University
of Southern California, Los Angeles, California 90089, USA
2Ming Hsieh Department of Electrical Engineering-Electrophysics, University of
Southern California, Los Angeles, California 90089, USA
*Corresponding author: firstname.lastname@example.org
Received July 25, 2013; revised September 26, 2013; accepted October 7, 2013;
posted October 8, 2013 (Doc. ID 194602); published November 11, 2013
The development of DNA analysis methods is rapidly expanding as interest in characterizing subtle variations in-
creases in biomedicine. A promising approach is based on evanescent field sensors that monitor the hybridization
with ssDNA using an epoxide method, and the evanescent wave of the microresonator excites a fluorescent label on
the complementary ssDNA during hybridization. Based on a temporal analysis, the different binding regimes can be
identified.© 2013 Optical Society of America
OCIS codes:(230.5750) Resonators; (130.6010) Sensors; (170.6280) Spectroscopy, fluorescence and luminescence.
As the genome is deciphered, the relationship between
subtle variations in DNA and various disease states is
becoming increasingly apparent [1–6]. To this end, re-
searchers are developing diagnostic platforms based
on DNA hybridization arrays [5–10]. DNA hybridization
or renaturation is the process in which a single strand
of nucleotides forms a noncovalent association with its
complement strand . The hybridization rate is gov-
erned by several parameters, including the degree of
similarity between the strands and environmental fac-
tors, such as pH and temperature.
contribute to hybridization, the measurement of DNA
pendent on the concentration of DNA, the temperature,
and the cation concentration. At the very minimum, there
are two conditions that must be met in order for hybridi-
zation to occur. For one, the salt concentration must be
are not repulsed. Second, the temperature of the sample
(Tm). In this temperature range, random intrastrand
hydrogen bonding is disrupted, favoring interstrand
hydrogen bonding between complementary base pairs.
The conventional method for studying hybridization is
to measure the reverse process, specifically the melting
temperature. In this approach, ssDNA is immobilized on
a surface and hybridized to the complement strand. The
sample is then gradually heated, denaturing or melting
the hybridized DNA, while recording the fluorescence
or optical absorbance spectra. From this signal, the bind-
ing energy can be determined. While this measurement
does provide an accurate reading of the dissociation of
the DNA, it is unable to yield information about the
behavior of DNA at ambient temperatures and physio-
logical pH and requires a significant quantity of the DNA.
Recently, researchers have recognized this challenge
and begun to develop new technologies to address it.
One approach is the combination of evanescent wave
(ssDNA-dye) [9,10]. In this approach, the ssDNA is immo-
bilized directly on the surface of the optical device
and exposed to the ssDNA-dye complement. The optical
field excites the dye molecules and the fluorescence is
detected using a variety of methods, including spectro-
graphs and power meters. Using this approach, research-
ers have shown the ability to detect the hybridization
process using these sensor devices.
process. Specifically, when molecules enter the evanes-
cent field, some will be oriented correctly and begin to
hybridize immediately; however, many will bind nonspe-
cifically to the device surface, rapidly dissociating and
diffusing away. These molecules will produce a transient
signal. Therefore, the detection signal has two compo-
nents: (1) the hybridized DNA and (2) nonhybridized
DNA. This two-phase process follows directly from mass
Clearly, during the initial phase of the experiment, the
second component (nonspecific binding) is significantly
larger than the first (hybridized DNA). However, as the
experiment progresses, it is anticipated that the signal
from the second component will go to zero, leaving only
the signal from the hybridized DNA. In previous work,
this dual-phase detection signal was not resolvable,
and possibly incorrectly interpreted as a single signal,
for several different reasons. For example, without suf-
ficiently high sensitivity and/or fast temporal resolution,
the transient signal would not be recorded by the detec-
tor (e.g., spectrograph). However, it is critical to fully
understand the temporal aspect of the sensor response,
given the time dependence of the DNA hybridization
In the present work, we use an evanescent wave
sensor in combination with a high speed spectrograph
to detect both phases of binding. The sensor is based
on an integrated optical resonant cavity, specifically a
toroidal microcavity [Fig. 1(a)] [13,14]. The 20-mer
ssDNA is attached to the surface of the device using
an epoxy approach, and its complement is labeled with
4690OPTICS LETTERS / Vol. 38, No. 22 / November 15, 2013
0146-9592/13/224690-04$15.00/0© 2013 Optical Society of America
the cyanine dye 5 (Cy5), which fluorescently emits be-
tween 650 and 670 nm when excited with a wavelength
around 635 nm. As the DNA hybridizes, the emission is
detected on the spectrograph and analyzed.
COMSOL Multiphysics finite element method modeling
of the optical field distribution inside the toroid and the
environment was performed to verify that the function-
alized DNA fell within the evanescent field of the toroid.
By leveraging the symmetry of the device, the modeling
challenge simplifies to a 2D problem, as shown in
Fig. 1(b). The evanescent field was found to extend ap-
proximately 100 nm into the environment, therefore,
completely overlapping the ssDNA and fluorophore.
Because of the long photon lifetime within the cavity
(high quality factor), very low input power is needed
to generate a strong fluorescent signal. In previous work,
optical cavities have demonstrated the ability to excite
fluorescent dyes embedded within lipid bilayers and
detect DNA [15–19].
The silica toroidal cavities are fabricated on silicon
wafers using a simple three-step method involving photo-
lithography, two etching processes (buffered oxide etch
and xenon difluoride etch), and a carbon dioxide laser
reflow step [13,14].
To immobilize and orient the ssDNA on the surface
of the cavity, we use an epoxide approach, which
relies on an amine-initiated nucleophilic ring-opening
reaction of an epoxide. Specifically, the surface of the
cavity is hydroxylated using an oxygen plasma (120 W,
200 mTorr, 30 sccm for 5 min). Then the epoxy linker
added by vapor deposition under vacuum for 45 min.
Finally, the amine modified oligonucleotides (IDT,
Coralville Iowa) are covalently linked to the epoxide
at 37°C in a humid environment for six hours or overnight
(Fig. 2) .
To verify the activity of the surface chemistry, a multi-
color fluorescent microscopy study is performed in
which the surface immobilized amine modified ssDNA
is labeled with 6-FAM, a derivative of fluorescein and
highly compatible with oligonucleotides, and the comple-
ment ssDNA is labeled with Cy5 (ssDNA-Cy5). As such,
6-FAM emits in the green and Cy5 emits in the red.
Imaging is performed at each step in the process: (1) after
hydroxylation and attachment of epoxide silane, (2) after
attachment of the amine modified, 6-FAM labeled
ssDNA, and (3) after hybridization of complement ssDNA
(ssDNA-Cy5) at room temperature. These reaction
conditions are identical to those used in the optical
device experiments and therefore serve as a control
measurement verifying hybridization. The images are
combined using Nikon NIS element basic research imag-
ing software. The presence of green fluorescence in
Fig. 3(b) indicates that this approach uniformly conju-
gates ssDNA to the surface of the device. Figure 3(c)
shows the emission from the ssDNA-Cy5 where it has
hybridized to the 6-FAM labeled ssDNA verifying that
hybridization occurs at room temperature. Figure 3(d)
is the overlap of these two images, showing precisely
where the two ssDNAs are colocated.
electron micrograph of a toroidal optical microcavity. (b) 2D
COMSOL finite element method simulation of the cross section
of a toroidal optical cavity. The optical field extends into the
environment, enabling excitation of fluorophores located near
the surface of the device.
Silica toroidal optical microcavity. (a) Scanning
cavity is hydroxylated and then GPTMS is used to covalently
attach epoxide groups. Animated ssDNA binds to the epoxides,
forming the ssDNA functionalized devices. In the last step, the
complement ssDNA-Cy5 is hybridized to the surface. Cy5 emits
in the red. To perform the control fluorescent imaging experi-
ments, the aminated ssDNA was labeled with 6-FAM, a green
Epoxide functionalization process. The surface of the
of a microtoroid after GPTMS vapor deposition and incubation
with amine modified ss-DNA-6-FAM. (b) Fluorescent image of
the same microtoroid showing the attachment of ss-DNA-
6-FAM. (c) Fluorescent image of a microtoroid after incubation
with complement ss-DNA-Cy5. The filters are adjusted to isolate
the Cy5 emission from the FAM emission. (d) Overlap of parts
(b) and (c), verifying that hybridization occurred. The specific
sequences used are: 3’-NH2-GCC GGA TAG CGT AAA GGT
TA-FAM and 5’-CGG CCT ACT GCA TTT CCA AT/Cy5-3’.
Multicolor fluorescent imaging. (a) Bright field image
November 15, 2013 / Vol. 38, No. 22 / OPTICS LETTERS4691
Two different optical measurements are performed:
(1) device characterization and (2) DNA hybridization
detection. For device characterization, we initially
immersed the bioconjugated resonate cavity in nuclease
free water and the DNA hybridization detection experi-
ments were measured in 100 μL of 2× sodium citrate
(SSC) with 0.2% sodium dodecyl sulfate (SDS) buffer
(pH 7.0) at ambient temperature. Light is coupled into
the device from a tunable narrow linewidth laser
centered at 633 nm (Velocity series, Newport) using a
tapered optical fiber waveguide.
To measure the photon lifetime (quality factor) of the
bioconjugated cavity, the cavity is aligned to the wave-
guide using a high precision three-axis nanopositioning
stage (Optosigma) and monitored on top and sideview
machine vision systems. The transmission spectrum
is recorded using a high-speed digitizer/oscilloscope
(National Instruments) and fit to a Lorentzian. The qual-
ity factor is calculated using the expression: Q ? λ∕δλ,
where δλ is the full width half-maximum of the Lorentzian
fit and λ is the resonant wavelength, resulting in a Q of
2.2 × 107for the device tested [Fig. 4(a)] [13,14].
To perform the hybridization detection experiments,
the testing setup is slightly modified, and a fiber-coupled
imaging spectrograph (Andor Shamrock SR-163 with a
Newton CCD detector) is integrated to the top [Fig. 4(b)]
. A filter that blocks light below 650 nm is placed
between the spectrograph tip and the toroid to avoid
saturation of the spectrograph detector with the laser
pump light. The complement ssDNA-Cy5 is injected into
the sample chamber using a syringe pump at a flow rate
of 50 μL∕min for two minutes. Cy5 was specifically
selected as the fluorophore because 633 nm falls within
the absorption range, allowing the evanescent field of the
cavity to efficiently excite the dye. The emission spectra
are continuously recorded for 30 minutes, and the peak
emission intensity of the dye (670 nm) is monitored as it
evolves throughout the experiment.
To verify and characterize the sensing method, two dif-
ferent detection measurements are performed using two
different functionalized devices. The first is detection of a
single solution of 2 μM ssDNA-Cy5. The second is the
characterization of the working range. For this measure-
ment, a series of ssDNA-Cy5 solutions are made with
concentrations ranging from 1 nM to 2 μM. The solutions
are then sequentially injected into the volume around the
toroidal cavity. Control experiments are also performed
without the ssDNA-Cy5 present.
Figure 5(a) shows a pair of representative excitation/
emission spectra. The excitation source at 633 nm, which
originates from the optical cavity, is clearly identifiable in
both spectra. However, the second peak, which corre-
sponds to the emission from the fluorescent dye, only
occurs when the ssDNA-Cy5 is present. The peak emis-
sion wavelength of the dye agrees with known values. It
is also important to note that the signal to noise ratio of
this measurement is extremely high.
Representative hybridization detection results are con-
tained in Fig. 5(b). There are two clearly identifiable
peaks. The first transient peak is very large in magnitude
but fades quickly. This peak is due to the ssDNA-Cy5
nonspecifically binding to the surface of the device or
diffusing through the evanescent field in solution. In
contrast, the second peak is smaller but stable, indicating
that the DNA has hybridized to the surface.
Figure 6 shows the characterization of the working
range of the device. As expected, it has a sigmoidal
spectra used to determine the quality factor of the cavity. Based
on the linewidth, this device has a Q of 2.2 × 107in water. (b) A
rendering of the testing setup, which can simultaneously record
the emission from the fluorescent dye using a fiber coupled
spectrograph and inject the ssDNA-Cy5 using a syringe pump.
The emission from the 633 nm tunable laser is partially blocked
using a red filter. The quality factor can also be measured in this
Optical device characterization. (a) A transmission
with and without the ssDNA-Cy5 present. While the 633 nm la-
ser line is present in both spectra, the fluorescent emission is
only present when the ssDNA-Cy5 is injected, as expected.
There are no secondary lines or other noise sources present
in this wavelength range. As such, the signal fidelity is
extremely high. The arrow indicates the 670 nm wavelength
which is tracked in the detection experiments. (b) The maxi-
mum of the emission at 670 nm, indicated in part (a), is moni-
tored and recorded while the ssDNA-Cy5 is injected. A strong
but transient signal is generated when the molecule nonspecifi-
cally binds to the surface and/or moves within the evanescent
field. The second stable peak is the result of the hybridization.
Detection of 2 μM ssDNA-Cy5. (a) Emission spectrum
exposed to several different ssDNA-Cy5 solutions.
Working range of the device as it is sequentially
4692OPTICS LETTERS / Vol. 38, No. 22 / November 15, 2013
portant to note that a significant amount of the potential
detection signal (Cy5 emission) is lost due to scattering
at the different interfaces and the optical absorption
ide based method for attaching ssDNA to the surface of
integrated optical cavities without degrading the optical
performance of the device. Using the evanescent field
of the cavity as an excitation source, we have detected
both nonspecific binding and the final hybridization
process. A more complete understanding of the various
transport processes that give rise to the detection signals
in DNA hybridization sensors as well as new diagnostic
methods will aid in developing improved biological
models for disease.
The authors would like to thank Soheil Soltani, Emma
Meinke, and Ce Shi at the University of Southern
California for assistance with FEM and ChemDraw. This
work was supported by the NIH New Innovators Award
Program (1DP2OD007391-01) and the Congressionally
Directed Medical Research Program (W81XWH-10-1-
1. D. F. Conrad, D. Pinto, R. Redon, L. Feuk, O. Gokcumen,
Y. J. Zhang, J. Aerts, T. D. Andrews, C. Barnes, P. Campbell,
T. Fitzgerald, M. Hu, C. H. Ihm, K. Kristiansson, D. G.
MacArthur, J. R. MacDonald, I. Onyiah, A. W. C. Pang, S.
Robson, K. Stirrups, A. Valsesia, K. Walter, J. Wei, C. Tyler-
Smith, N. P. Carter, C. Lee, S. W. Scherer, and M. E. Hurles,
Nature 464, 704 (2009).
2. D. Hanahan and R. A. Weinberg, Cell 144, 646 (2011).
3. S. P. Jackson and J. Bartek, Nature 461, 1071 (2009).
4. R. Lister, M. Pelizzola, R. H. Dowen, R. D. Hawkins, G. Hon,
J. Tonti-Filippini, J. R. Nery, L. Lee, Z. Ye, Q. M. Ngo,
L. Edsall, J. Antosiewicz-Bourget, R. Stewart, V. Ruotti,
A. H. Millar, J. A. Thomson, B. Ren, and J. R. Ecker, Nature
462, 315 (2009).
5. A. Lujambio, G. A. Calin, A. Villanueva, S. Ropero, M.
Sanchez-Cespedes, D. Blanco, L. M. Montuenga, S. Rossi,
M. S. Nicoloso, W. J. Faller, W. M. Gallagher, S. A. Eccles,
C. M. Croce, and M. Esteller, Proc. Natl. Acad. Sci. USA
105, 13556 (2008).
6. L. Zhang, J. Huang, N. Yang, J. Greshock, M. S. Megraw,
A. Giannakakis, S. Liang, T. L. Naylor, A. Barchetti, M. R.
Ward, G. Yao, A. Medina, A. O’Brien-Jenkins, D. Katsaros,
A. Hatzigeorgiou, P. A. Gimotty, B. L. Weber, and G.
Coukos, Proc. Natl. Acad. Sci. USA 103, 9136 (2006).
7. R. B. Stoughton, Annu. Rev. Biochem. 74, 53 (2005).
8. D. J. Lockhart, H. L. Dong, M. C. Byrne, M. T. Follettie, M. V.
Horton, and E. L. Brown, Nat. Biotechnol. 14, 1675 (1996).
9. F. Long, S. Wu, M. He, T. Tong, and H. Shi, Biosens. Bioelec-
tron. 26, 2390 (2011).
10. L. Malic, T. Veres, and M. Tabrizian, Biosens. Bioelectron.
26, 2053 (2011).
11. P. S. Thomas, Proc. Natl. Acad. Sci. USA 77, 5201 (1980).
Nature 421, 925 (2003).
14. X. Zhang, H.-S. Choi, and A. M. Armani, Appl. Phys. Lett. 96,
15. L. M. Freeman and A. M. Armani, IEEE J. Sel. Top. Quantum
Electron. 18, 1160 (2012).
16. L. M. Freeman, S. Li, Y. Dayani, H. S. Choi, N. Malmstadt,
and A. M. Armani, Appl. Phys. Lett. 98, 143703 (2011).
17. H. K. Hunt and A. M. Armani, Nanoscale 2, 1544 (2010).
18. M. S. Luchansky and R. C. Bailey, Anal. Chem. 84, 793
Biophys. J. 85, 1974 (2003).
20. A. Carré, W. Birch, and V. LaCarriére, Silanes and
Coupling Agents, K. L. Mittal, ed. (Brill, 2007), p. 14.
November 15, 2013 / Vol. 38, No. 22 / OPTICS LETTERS 4693