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Optics Express

Optics Express

  • Editor: C. Martijn de Sterke
  • Vol. 17, Iss. 26 — Dec. 21, 2009
  • pp: 24224–24233
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Sub-wavelength nanofluidics in photonic crystal sensors

Min Huang, Ahmet Ali Yanik, Tsung-Yao Chang, and Hatice Altug  »View Author Affiliations


Optics Express, Vol. 17, Issue 26, pp. 24224-24233 (2009)
http://dx.doi.org/10.1364/OE.17.024224


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Abstract

We introduce a novel sensor scheme combining nano-photonics and nano-fluidics on a single platform through the use of free-standing photonic crystals. By harnessing nano-scale openings, we theoretically and experimentally demonstrate that both fluidics and light can be manipulated at sub-wavelength scales. Compared to the conventional fluidic channels, we actively steer the convective flow through the nanohole openings for effective delivery of the analytes to the sensor surface. We apply our method to detect refractive index changes in aqueous solutions. Bulk measurements indicate that active delivery of the convective flow results in better sensitivities. The sensitivity of the sensor reaches 510 nm/RIU for resonance located around 850 nm with a line-width of ~10 nm in solution. Experimental results are matched very well with numerical simulations. We also show that cross-polarization measurements can be employed to further improve the detection limit by increasing the signal-to-noise ratio.

© 2009 OSA

1. Introduction

In recent years, label free bio-sensors combined with innovative signal transduction methods are proposed to push the detection limits down to femto-molar concentrations of analytes [1

1. A. N. Shipway, E. Katz, and I. Willner, “Nanoparticle arrays on surfaces for electronic, optical, and sensor applications,” ChemPhysChem 1(1), 18–52 (2000). [CrossRef]

3

3. D. Erickson, S. Manda, H. J. Allen, Yang, and B. Cordovez, “Nanobiosensors: optofluidic, electrical and mechanical approaches to biomolecular detection at the nanoscale.” Microfluid. Nanofluid. 4(1–2), 33–52 (2007). [CrossRef]

]. Concurrently, researchers are integrating such sensitive and compact nano-sensors with micro-fluidics for automated sample handling [4

4. P. S. Waggoner and H. G. Craighead, “Micro- and nanomechanical sensors for environmental, chemical, and biological detection,” Lab Chip, Volume 7(10), 1238–1255 (2007). [CrossRef]

, 5

5. C. Monat, P. Domachuk, and B. J. Eggleton, “Integrated optofluidics: A new river of light,” Nature Photon. 1(2), 106–114 (2007). [CrossRef]

]. While micro-fluidics can enable portable and lab-on-a-chip systems, recent theoretical and numerical calculations indicate that we have to take into account the effects of various fluidic integration schemes as they can fundamentally limit the sensor performances [6

6. B. Kuswandi, J. Nuriman, J. Huskens, and W. Verboom, “Optical sensing systems for microfluidic devices: A review,” Analytica Chimica Acta 601(2), 141–155 (2007). [CrossRef] [PubMed]

]. For nano-sensors embedded in conventional microfluidic channels, the detection limit is often determined by the analyte (mass) transport limitations as opposed to the detection capabilities of the sensors [7

7. P. E. Sheehan and L. J. Whitman, “Detection limits for nanoscale biosensors,” Nano Lett. 5(4), 803–807 (2005). [CrossRef] [PubMed]

]. As the analytes are collected by the functionalized sensors, depletion zones form around the sensing area. Depletion zone, where the analytes transport diffusively, expands with time until its growth is halted by the convective flow [8

8. J. Bishop, S. Blair, and A. Chagovetz, “Convective flow effects on DNA biosensors,” Biosens. Bioelectron. 22(9-10), 2192–2198 (2007). [CrossRef]

, 9

9. T. M. Squires, R. J. Messinger, and S. R. Manalis, “Making it stick: convection, reaction and diffusion in surface-based biosensors,” Nature Biotechnol. 26(4), 417–426 (2008). [CrossRef]

]. In micro-fluidic channels supporting laminar flow profile, the convective flow parallel to the surface is weaker close to the channel edge. Accordingly, the depletion zones extend significantly towards the center of the channel. It causes dramatically lower amounts of analytes to reach the sensing surface per unit time [10

10. J. P. Golden, T. M. Floyd-Smith, D. R. Mott, and F. S. Ligler, “Target delivery in a microfluidic immunosensor,” Biosens. Bioelectron. 22(11Issue 11), 2763–2767 (2007). [CrossRef] [PubMed]

]. Consequently, if no method is introduced to actively direct the convective flow towards the surface of the nano-micro size sensors, analytes at low concentrations may need week-to-years to diffuse due to mass (analyte) transport limitations imposed by the depletion zones [7

7. P. E. Sheehan and L. J. Whitman, “Detection limits for nanoscale biosensors,” Nano Lett. 5(4), 803–807 (2005). [CrossRef] [PubMed]

].

2. Device scheme and the flow analysis

Figure 1(c) and (d) show the steady state velocity distribution for the actively (proposed) and the passively (conventional) controlled convective flow schemes. Flow profiles around PhC regions are shown in detail (insets). For the passively controlled scheme (Fig. 1(d)), as the viscous forces in the fluid dominate over the inertial forces, we observe the formation of laminar flow profile. The convective flow is fast close to the center of the channel but becomes very slow near the edges. This implies that in an immunoassay based sensing the depletion zones will extend further from the sensor surface causing ever slower analyte transport. One can increase the convective flow rate to shrink the depletion zones. However, such a passive (indirect) control only results in moderate improvements in mass transport rates [9

9. T. M. Squires, R. J. Messinger, and S. R. Manalis, “Making it stick: convection, reaction and diffusion in surface-based biosensors,” Nature Biotechnol. 26(4), 417–426 (2008). [CrossRef]

]. The alternative approach we propose here to overcome the mass transport limitation is to steer the convective flow directly towards the sensing surface. This is demonstrated in microfludic simulation in Fig. 1(c) where the convective flow is still very strong around the sensing surface and the turbulences (stirring of the solution) are generated around the holes. Such a directed flow can strongly improve the delivery of the analytes to the sensor surface. This scheme also helps to overcome the surface tension of highly viscous solution and guarantees that the sensor can be totally immersed in solution. In this way, as both sides of the structure are exposed to the solution, the sensitivity is further enhanced.

3. Device fabrication and sensor design

3.1 Device fabrication

In order to implement the proposed scheme, we use PhC structures on free standing membranes. One important consideration here is the mechanical strength of the membranes as they need to stand the relatively high pressures generated by the perpendicular convective flow. Mechanically highly robust Low Pressure Chemical Vapor Deposition (LPCVD) silicon nitride (SiNx) films are excellent choice. In addition, LPCVD SiNx films have very good optical properties for the implementation of PhCs. They are transparent in the visible/near-infrared regime with high refractive index. Figure 2(a)
Fig. 2 (a) Fabrication scheme. Structure is first patterned on Polymethyl methacrylate (PMMA) layer using EBL. Then reactive ion etching (RIE) is used to dry etch the SiNx slab. PMMA left on the structure is cleaned using O2 plasma asher, resulting in a suspended PhC membrane. (b)-(c) SEM top view of the structure. (d) SEM cross-view of the structure titled at 40°.
summarizes the fabrication steps. We start with ~200 x 200 μm2 area free-standing SiNx membranes. PhCs are fabricated by performing e-beam lithography (EBL) and dry-etching. Scanning Electron Microscopy (SEM) images of the fabricated structures are shown in Fig. 2(b), 2(c) and 2(d). The tilted SEM image taken at 40° confirms the vertical wall profiles of the openings. The thickness of the fabricated membranes (~90 nm) is much smaller compared to the membrane area (~40,000 μm2). Although the aspect ratio is very high, the membranes are observed to survive long hours of operation under flow pressure.

3.2 Photonic crystal sensor design

PhCs offer unique opportunities to tailor the spatial extent of the electromagnetic field and control the strength of the light-matter interaction. In this work, we exploit guided resonances that are delocalized in the plane and tightly confined in the vertical direction. The periodic index contrast of the structures enables the excitation of the guided resonances with a plane-wave illumination at normal incidence and their out-coupling into the radiation modes. Such a surface normal operation eliminates the alignments of sensitive prism/waveguide/fiber coupling schemes needed by other optical nanosensors [21

21. D. Nedelkov and R. W. Nelson, “Surface plasmon resonance mass spectrometry: recent progress and outlooks,” Trends in Biotechnology, Volume 21(7Issue 7), 301–305 (2003). [CrossRef]

24

24. S. Chana, Y. Li, L. J. Rothberg, B. L. Miller, and P. M. Fauchet, “Nanoscale silicon microcavities for biosensing.” Mat. Scie. Engin. C 15(1–2), 277–282 (2001). [CrossRef]

]. The ease of resonance excitation by surface normal light is particularly advantageous for high-throughput micro-array applications. The incident light is transmitted by PhC slabs through two different pathways [25

25. S. Fan and J. D. Joannopoulos, “Analysis of guided resonances in photonic crystal slabs,” Phys. Rev. B 65(23), 235112 (2002). [CrossRef]

]. One of them is the direct pathway, where a portion of the electromagnetic field goes straight through the slab. The other is the indirect pathway, where the remaining portion couples into the guided resonances before leaking into the radiation modes. These two pathways interfere with each other and result in resonances with sharp Fano-type asymmetric line-shapes. The spectral location of these resonances is highly sensitive to the refractive index changes occurring within the surroundings of PhC slabs. The index change due to the accumulation of bio-molecules or variations in the bulk solution could be detected optically in a label-free fashion.

To experimentally implement the proposed sensor, we use square lattice SiNx PhC slabs (inset in Fig. 3(a)
Fig. 3 PhC sensor design: (a) Transmission spectra calculated by 3D-FDTD simulations are shown when the PhC slab is emerged in three different media: air (blue), water (red) and an IPA-chloroform mixture (green), respectively. Inset shows the schematic view of the design. Parameters for the structure are: r = 270 nm, a = 600 nm and d=90nm. (b) Electromagnetic intensity distribution of the 1st mode when the structure is in air. Top and cross section views are shown, respectively.
). Figure 3(a) shows the transmission spectra of a specific design calculated by three dimensional finite-difference time-domain (3D-FDTD) method in three different media: air (refractive index n = 1), water (n = 1.33), and an IPA-chloroform mixture (n = 1.43). A normally incident plane wave source (corresponding to the Γ-point in the dispersion diagram) excites the eigenmodes of the system. For each case, two modes are observed within the given spectral range. Figure 3(b) shows the intensity distribution of the lowest (first) order mode when the structure is in air. The field has four-fold symmetry as the lattice and well confined within the slab in the vertical direction. Within the plane, the field extends into the holes, which is crucial in increasing the field overlap with the surrounding media for higher sensitivity. We evaluate its bulk sensitivity (in units of nm/RIU) by calculating the shift of the resonance position in wavelength versus the refractive index change in the surrounding environment. To optimize the structure for higher sensitivity, we studied the effects of the slab thickness and the hole radius by varying the thickness d from 0.1a to 0.3a and the radius r from 0.3a to 0.45a (a is the periodicity). For all the analyzed structures, the resonant wavelength of the lowest order mode in air is scaled to 670 nm. The calculated sensitivities and the parameter sets for each case are given in Table 1

Table 1. Sensitivity results with different hole radius and slab thickness (in unit of nm/RIU)

table-icon
View This Table
. The sensitivity improves as the size of the holes increases and the slab thickness decreases. When r=0.45a and d=0.1a, the sensitivity reaches 560 nm/RIU. Given the difficulty of fabricating such a thin membrane, we instead use the design with thickness d=0.15a. Its sensitivity is still over 535 nm/RIU. As the sensitivity scales with wavelength, shifting the resonances to the longer wavelength (such as 1550 nm range) could increase the sensitivity even further (well above 1000 nm/RIU).

The optimized PhC structures are fabricated on free standing SiNx membranes according to the process flow described in Fig. 2. SEM images indicate that the diameter and the periodicity are 540 nm and 605 nm, respectively. Ellipsometer measurements are taken on the unpatterned area of the membrane to confirm that the slab thickness is ~90 nm. These numbers are quite close to the optimized design with r/a=0.45 and d/a=0.15. For the PhC with periodicity of 605 nm, the resonance peak in air is located at ~670nm. This wavelength is well within the spectral coverage of our experimental setup.

4. Experimental results

4.1 Controlling the flow

To carry out the flow tests, the structures are integrated in a chamber with two inlets/outlets both on the top and the bottom channels fabricated in polydimethylsiloxane (PDMS). To implement the laminar flow scheme, where the convective flow is parallel to the surface (Fig. 1(b) and Fig. 1(d)), we blocked the inlet/outlet of the bottom channel. To steer the convective flow actively towards the sensing surface, we blocked one of the openings of the both channels (Fig. 1(a) and Fig. 1(c)). The PhC slab is sealed perfectly to ensure the flow is only through the openings. Video images of the perpendicular convective flow, captured in a microscope with a CCD camera, are shown in Fig. 4
Fig. 4 (Media 1) images of the actively directed perpendicular convective flow: (a) Bottom channel is almost filled up with IPA. (b) IPA starts to go through the nanohole openings. (c) IPA spreads over the surface. (d) The whole structure is emerged in IPA.
. (video stream is available in the supplementary). Here, the IPA solution is pumped into the bottom channel by a syringe at a rate of 80μL/s. The video recording starts when the bottom channel is almost filled-up. Figure 4 (and video) shows the merge of IPA to the top channel only through the openings, confirming the active steering of the liquid flow. No damage or breakage of the membrane due to the applied pressure is observed.

4.2 Sensor response comparison for the actively controlled and conventional schemes

4.3 Sensor sensitivity for the actively controlled delivery scheme

Bulk sensitivity of the PhCs is tested by successively applying five different solutions through the directed flow scheme: DI-water, acetone, IPA and two IPA-chloroform mixtures with refractive indices of 1, 1.33, 1.356, 1.377, 1.401 and 1.424, respectively. The refractive indices of all the liquids are initially measured using a commercial refractometer. The measurements are performed by slowly pumping the solution to the chamber at 50 μL/s pumping rate. Prior to each measurement, we make sure the former solution is entirely replaced by the new one. As shown in Fig. 6(a)
Fig. 6 (a) Experimentally measured transmission spectra of PhC slab using actively controlled delivery scheme in air (blue), water (red), IPA (green) and an IPA-chloroform mixture (black). (b) Shifts of the 1st resonant peaks in wavelength versus the surrounding refractive index. Resonance peak positions found in experiments (blue stars) match very well with the simulation results (green circles). Red line is a linear fitting to the experimental results.
, with increasing refractive index the resonances red-shift and the line-widths become narrower. The linewidth of the resonance in DI-water is measured to be ~10 nm. Figure 6(b) shows the shift in resonance wavelength versus the refractive index of the liquid. The agreement between the experimental data and the theoretically predicted shifts is excellent. The experimentally measured sensitivity of the sensor, 510 nm/RIU for operation near 850 nm in wavelength, is much larger than the previously reported values [20

20. O. Levi, M. M. Lee, J. Zhang, V. Lousse, S. R. J. Brueck, S. Fan, and J. S. Harris, “Sensitivity analysis of a photonic crystal structure for index-of-refraction sensing,” Proc. SPIE 6447, 2–9 (2007).

, 26

26. I. D. Block, N. Ganesh, M. Lu, and B. T. Cunningham, “A sensitivity model for predicting photonic crystal biosensor performance,” IEEE Sensors 8(3), 274–280 (2008). [CrossRef]

].

4.4 Isolating the resonances with large signal-to-noise ratio

As we show above, with the demonstrated sensor system, we can effectively detect the refractive index changes by tracking the resonances. However, minute amounts of analytes from small quantities of biological samples would result in very small resonance peak shifts. In such cases, it is crucial to have narrow resonances with large signal-to-noise ratios. This can be achieved by using cross-polarization measurements [17

17. H. Altug and J. Vuckovic, “Polarization control and sensing with two-dimensional coupled photonic crystal microcavity arrays,” Opt. Lett. 30(9), 982 (2005). [CrossRef] [PubMed]

,27

27. K. A. Tetz, L. Pang, and Y. Fainman, “High-resolution surface plasmon resonance sensor based on linewidth-optimized nanohole array transmittance,” Opt. Lett. 31(10), (1528–1530).

]. As mentioned above, the transmission spectra result from interference of two optical paths: one is the direct transmission while the other is through the guided resonances. When we employ an unpolarized light and collect all the light transmitted through the slab, both pathways contributes to the detected signal. However, if we launch a polarized light and collect the signal after an analyzer oriented perpendicular to the polarizer, only the scattering from the guided resonances contributes. This results in dramatic suppression of the background and isolation of the resonances with large signal-to-noise ratios. In addition, the cross-polarization measurements result in purely Lorentzian-shape resonance profiles with narrower line-widths. Figure 7(a)
Fig. 7 (a) Cross polarization spectrum (blue) and the regular unpolarized measurement (red). (b) Green curves correspond to the fitting of the resonance feature in the spectrum with single Lorentzian functions. Their summation is denoted in red dashed line and overlaid with the experimental result (blue curve).
compares the cross-polarization spectrum (red) with the regular one (blue). The spectra are taken when the structure is in air. Cross-polarization measurements clearly isolate two distinct resonance features from the background. A single Lorentzian with 7 nm line-width fits very well with the second order mode resonance (Fig. 7(b)). On the other hand, two Lorenztians are needed to fit the lowest order mode (Fig. 7(b)). This indicates a potential resonance splitting for the lowest order mode, which could be due to a slight non-uniformity in fabrication. The addition of three Lorentzians (red dashed curve in Fig. 7(b)) matches very well with the experimentally measured spectrum.

5. Conclusion

In conclusion, we introduced a novel sensor combining nano-photonics and nano-fluidics on a single platform. By using nano-scale openings in PhCs, we show that both light and fluidics can be manipulated on chip. We present both theoretical and experimental analysis of the fluidic and the photonic components of the integrated system. Compared to the laminar flow in conventional fluidic channels, we show that active steering of the convective flow results in the direct delivery of the stream to the nanohole openings. This can lead to enhanced analyte delivery to the sensor surface by overcoming the mass transport limitations. We apply our method to detect refractive index changes in aqueous solutions. Bulk measurements show that actively directed convective flow results in better sensitivities. Experimental results are matched very well with the simulations. The sensitivity of the sensor reaches 510 nm/RIU for resonance located around 850 nm with a line-width of ~10 nm in solution. We show that cross-polarization measurement can be employed to further improve the detection limit by increasing the signal-to-noise ratio.

Acknowledgment

Authors thank Rui Zhang for discussions. This work is supported in part by NSF SGER Award (ECSS-0849603), Massachusetts Life Science Center New Investigator Award (H.A.), Boston University Photonics Center and NSF Engineering Research Center on Smart Lighting (EEC-0812056).

References and links

1.

A. N. Shipway, E. Katz, and I. Willner, “Nanoparticle arrays on surfaces for electronic, optical, and sensor applications,” ChemPhysChem 1(1), 18–52 (2000). [CrossRef]

2.

R. Raiteria, M. Grattarola, and R. Berge, “Micromechanics senses biomolecules,” Materials Today 5(1), 22–29 (2002). [CrossRef]

3.

D. Erickson, S. Manda, H. J. Allen, Yang, and B. Cordovez, “Nanobiosensors: optofluidic, electrical and mechanical approaches to biomolecular detection at the nanoscale.” Microfluid. Nanofluid. 4(1–2), 33–52 (2007). [CrossRef]

4.

P. S. Waggoner and H. G. Craighead, “Micro- and nanomechanical sensors for environmental, chemical, and biological detection,” Lab Chip, Volume 7(10), 1238–1255 (2007). [CrossRef]

5.

C. Monat, P. Domachuk, and B. J. Eggleton, “Integrated optofluidics: A new river of light,” Nature Photon. 1(2), 106–114 (2007). [CrossRef]

6.

B. Kuswandi, J. Nuriman, J. Huskens, and W. Verboom, “Optical sensing systems for microfluidic devices: A review,” Analytica Chimica Acta 601(2), 141–155 (2007). [CrossRef] [PubMed]

7.

P. E. Sheehan and L. J. Whitman, “Detection limits for nanoscale biosensors,” Nano Lett. 5(4), 803–807 (2005). [CrossRef] [PubMed]

8.

J. Bishop, S. Blair, and A. Chagovetz, “Convective flow effects on DNA biosensors,” Biosens. Bioelectron. 22(9-10), 2192–2198 (2007). [CrossRef]

9.

T. M. Squires, R. J. Messinger, and S. R. Manalis, “Making it stick: convection, reaction and diffusion in surface-based biosensors,” Nature Biotechnol. 26(4), 417–426 (2008). [CrossRef]

10.

J. P. Golden, T. M. Floyd-Smith, D. R. Mott, and F. S. Ligler, “Target delivery in a microfluidic immunosensor,” Biosens. Bioelectron. 22(11Issue 11), 2763–2767 (2007). [CrossRef] [PubMed]

11.

R. W. Boyd and J. E. Heebner, “Sensitive disk resonator photonic biosensor,” Applied Optics, Volume 40(31Issue 31), 5742–5747 (2001). [CrossRef]

12.

A. J. Haes, S. Zou, G. C. Schatz, and R. P. Van Duyne, “A Nanoscale Optical Biosensor: The Long Range Distance Dependence of the Localized Surface Plasmon Resonance of Noble Metal Nanoparticles,” J. Phys. Chem. B 108(1), 109–116 (2004). [CrossRef]

13.

A. J. Haes and R. P. Van Duyne, “A unified view of propagating and localized surface plasmon resonance biosensors.” Anal. Bioanal. Chem. 379(7–8), 920–930 (2004). [CrossRef] [PubMed]

14.

A. D. Leebeeck, L. K. Swaroop Kumar, V. D. Lange, D. Sinton, R. Gordon, and A. G. Brolo, “On-Chip Surface-Based Detection with Nanohole Arrays,” Anal. Chem. 79(11), 4094–4100 (2007). [CrossRef] [PubMed]

15.

A. Artar, A. A. Yanik, and H. Altug, “Fabry–Pérot nanocavities in multilayered plasmonic crystals for enhanced biosensing,” Appl. Phy. Lett. 95(5), 051105 (2009). [CrossRef]

16.

E. Chow, A. Grot, L. W. Mirkarimi, M. Sigalas, and G. Girolami, “Ultracompact biochemical sensor built with two-dimensional photonic crystal microcavity,” Opt. Lett. 29(10), 1093–1095 (2004). [CrossRef] [PubMed]

17.

H. Altug and J. Vuckovic, “Polarization control and sensing with two-dimensional coupled photonic crystal microcavity arrays,” Opt. Lett. 30(9), 982 (2005). [CrossRef] [PubMed]

18.

N. Skivesen, A. Têtu, M. Kristensen, J. Kjems, L. H. Frandsen, and P. I. Borel, “Photonic-crystal waveguide biosensor,” Opt. Express 15(6), 3169–3176 (2007). [CrossRef] [PubMed]

19.

L. Shi, P. Pottier, Y.-A. Peter, and M. Skorobogatiy, “Guided-mode resonance photonic crystal slab sensors based on bead monolayer geometry,” Opt. Express 16(22), 17962–17971 (2008). [CrossRef] [PubMed]

20.

O. Levi, M. M. Lee, J. Zhang, V. Lousse, S. R. J. Brueck, S. Fan, and J. S. Harris, “Sensitivity analysis of a photonic crystal structure for index-of-refraction sensing,” Proc. SPIE 6447, 2–9 (2007).

21.

D. Nedelkov and R. W. Nelson, “Surface plasmon resonance mass spectrometry: recent progress and outlooks,” Trends in Biotechnology, Volume 21(7Issue 7), 301–305 (2003). [CrossRef]

22.

S. Mandal and D. Erickson, “Nanoscale optofluidic sensor arrays,” Opt. Express 16(3), 1623–1631 (2008). [CrossRef] [PubMed]

23.

A. M. Armani, R. P. Kulkarni, S. E. Fraser, R. C. Flagan, and K. J. Vahala, “Label-Free, Single-Molecule Detection with Optical Microcavities.” Science 317(5839), 783–787 (2007). [CrossRef] [PubMed]

24.

S. Chana, Y. Li, L. J. Rothberg, B. L. Miller, and P. M. Fauchet, “Nanoscale silicon microcavities for biosensing.” Mat. Scie. Engin. C 15(1–2), 277–282 (2001). [CrossRef]

25.

S. Fan and J. D. Joannopoulos, “Analysis of guided resonances in photonic crystal slabs,” Phys. Rev. B 65(23), 235112 (2002). [CrossRef]

26.

I. D. Block, N. Ganesh, M. Lu, and B. T. Cunningham, “A sensitivity model for predicting photonic crystal biosensor performance,” IEEE Sensors 8(3), 274–280 (2008). [CrossRef]

27.

K. A. Tetz, L. Pang, and Y. Fainman, “High-resolution surface plasmon resonance sensor based on linewidth-optimized nanohole array transmittance,” Opt. Lett. 31(10), (1528–1530).

OCIS Codes
(170.0170) Medical optics and biotechnology : Medical optics and biotechnology
(260.5740) Physical optics : Resonance
(280.1415) Remote sensing and sensors : Biological sensing and sensors
(350.4238) Other areas of optics : Nanophotonics and photonic crystals
(220.4241) Optical design and fabrication : Nanostructure fabrication
(050.6624) Diffraction and gratings : Subwavelength structures

ToC Category:
Photonic Crystals

History
Original Manuscript: October 21, 2009
Revised Manuscript: November 9, 2009
Manuscript Accepted: November 9, 2009
Published: December 18, 2009

Virtual Issues
Vol. 5, Iss. 1 Virtual Journal for Biomedical Optics

Citation
Min Huang, Ahmet Ali Yanik, Tsung-Yao Chang, and Hatice Altug, "Sub-wavelength nanofluidics in photonic crystal sensors," Opt. Express 17, 24224-24233 (2009)
http://www.opticsinfobase.org/oe/abstract.cfm?URI=oe-17-26-24224


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References

  1. A. N. Shipway, E. Katz, and I. Willner, “Nanoparticle arrays on surfaces for electronic, optical, and sensor applications,” ChemPhysChem 1(1), 18–52 (2000). [CrossRef]
  2. R. Raiteria, M. Grattarola, and R. Berge, “Micromechanics senses biomolecules,” Materials Today 5(1), 22–29 (2002). [CrossRef]
  3. D. Erickson, S. Manda, H. J. Allen, Yang, and B. Cordovez, “Nanobiosensors: optofluidic, electrical and mechanical approaches to biomolecular detection at the nanoscale.” Microfluid. Nanofluid. 4(1–2), 33–52 (2007). [CrossRef]
  4. P. S. Waggoner and H. G. Craighead, “Micro- and nanomechanical sensors for environmental, chemical, and biological detection,” Lab Chip, Volume 7(10), 1238–1255 (2007). [CrossRef]
  5. C. Monat, P. Domachuk, and B. J. Eggleton, “Integrated optofluidics: A new river of light,” Nature Photon. 1(2), 106–114 (2007). [CrossRef]
  6. B. Kuswandi, J. Nuriman, J. Huskens, and W. Verboom, “Optical sensing systems for microfluidic devices: A review,” Analytica Chimica Acta 601(2), 141–155 (2007). [CrossRef] [PubMed]
  7. P. E. Sheehan and L. J. Whitman, “Detection limits for nanoscale biosensors,” Nano Lett. 5(4), 803–807 (2005). [CrossRef] [PubMed]
  8. J. Bishop, S. Blair, and A. Chagovetz, “Convective flow effects on DNA biosensors,” Biosens. Bioelectron. 22(9-10), 2192–2198 (2007). [CrossRef]
  9. T. M. Squires, R. J. Messinger, and S. R. Manalis, “Making it stick: convection, reaction and diffusion in surface-based biosensors,” Nature Biotechnol. 26(4), 417–426 (2008). [CrossRef]
  10. J. P. Golden, T. M. Floyd-Smith, D. R. Mott, and F. S. Ligler, “Target delivery in a microfluidic immunosensor,” Biosens. Bioelectron. 22(11Issue 11), 2763–2767 (2007). [CrossRef] [PubMed]
  11. R. W. Boyd and J. E. Heebner, “Sensitive disk resonator photonic biosensor,” Applied Optics, Volume 40(31Issue 31), 5742–5747 (2001). [CrossRef]
  12. A. J. Haes, S. Zou, G. C. Schatz, and R. P. Van Duyne, “A Nanoscale Optical Biosensor: The Long Range Distance Dependence of the Localized Surface Plasmon Resonance of Noble Metal Nanoparticles,” J. Phys. Chem. B 108(1), 109–116 (2004). [CrossRef]
  13. A. J. Haes and R. P. Van Duyne, “A unified view of propagating and localized surface plasmon resonance biosensors.” Anal. Bioanal. Chem. 379(7–8), 920–930 (2004). [CrossRef] [PubMed]
  14. A. D. Leebeeck, L. K. Swaroop Kumar, V. D. Lange, D. Sinton, R. Gordon, and A. G. Brolo, “On-Chip Surface-Based Detection with Nanohole Arrays,” Anal. Chem. 79(11), 4094–4100 (2007). [CrossRef] [PubMed]
  15. A. Artar, A. A. Yanik, and H. Altug, “Fabry–Pérot nanocavities in multilayered plasmonic crystals for enhanced biosensing,” Appl. Phy. Lett. 95(5), 051105 (2009). [CrossRef]
  16. E. Chow, A. Grot, L. W. Mirkarimi, M. Sigalas, and G. Girolami, “Ultracompact biochemical sensor built with two-dimensional photonic crystal microcavity,” Opt. Lett. 29(10), 1093–1095 (2004). [CrossRef] [PubMed]
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