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Optics Express

  • Vol. 17, Iss. 7 — Mar. 30, 2009
  • pp: 5794–5806
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Retention of polarization signatures in SHG microscopy of scattering tissues through optical clearing

Oleg Nadiarnykh and Paul J. Campagnola  »View Author Affiliations


Optics Express, Vol. 17, Issue 7, pp. 5794-5806 (2009)
http://dx.doi.org/10.1364/OE.17.005794


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Abstract

Polarization responses in Second Harmonic Generation (SHG) imaging microscopy are a valuable method to quantify aspects of tissue structure, and may be a means to differentiate normal and diseased tissues. Due to multiple scattering, the polarization data is lost in turbid tissues. Here we investigate if this information can be retained through the use of optical clearing which greatly reduces the scattering coefficient and increases the corresponding mean free path. To this end, we have measured the SHG intensity as a function of laser polarization and the SHG signal anisotropy in murine tendon and striated muscle over a depth range of 200 microns. We find that the laser polarization is highly randomized in the uncleared tissues at depths corresponding to only 2-3 scattering collisions (50-10 microns). This depolarization of the laser is also reflected in the randomized anisotropy of the SHG signal as it is created over a range of polarization states. In strong contrast, both polarization signatures are significantly retained through ~200 microns of tissue thickness following treatment with 50% glycerol. Moreover, the measured polarization responses for both tendon and striated muscle are consistent with the extent of reduction of the respective scattering coefficients upon clearing. We suggest the method will be applicable to SHG imaging of connective disorders as well as cancer through several hundred microns of extracellular matrix.

© 2009 Optical Society of America

1. Introduction

Determining the health of tissues by in vivo imaging could impact diagnosis and monitoring of many diseases in a non-invasive or minimally invasive manner. However, currently used clinical technologies are hampered by insufficient resolution, lack of specificity, or high degree of invasiveness. While significant technological advances have greatly improved the capabilities of ultrasound, MRI, CT, and PET imaging these modalities are practically limited to resolutions of ~ 1 mm. Similarly diffuse optical tomography and photoacoustic tomography are limited to resolutions of ~ 1 mm and 100 microns, respectively. However, much higher spatial resolution (1 micrometer or less) is required to visualize structural changes associated with diseased cells and tissues. For example, it is increasingly recognized that changes in the microscopic tissue structure, e.g. the collagen fibril assembly in and around tumors, may also be early indicators of disease. To meet this need, we have been implementing Second Harmonic Generation (SHG) microscopy, which visualizes tissue assembly with submicron resolution, as a tool to differentiate normal and diseased states, (e.g. connective tissue disorders or cancers). For example, the type I collagen is abnormally assembled in osteogenesis imperfecta (brittle bone disease) and we found SHG could differentiate diverse tissues such as bone, skin, and tendon using metrics based on intensity comparisons, polarization analysis, and emission directionality.1, 2 We further showed statistical differentiation in several murine models of muscle disorders based on changes in sarcomere length.3 SHG has also shown early promise in cancer imaging as a stand alone technique,4-6 and it also has been implemented in multimodal approaches combining multiphoton excited fluorescence.7, 8 Given these successes, minimally invasive and truly in-vivo visualization of tissues may well progress into clinical techniques for assessing disease severity, and monitoring efficacy of treatment.

However, the increased resolution and specificity of multiphoton microscopy techniques comes at the expense of limited imageable penetration depths relative to established clinical modalities which typically image through several mm or cms of tissue. As opposed to this diffuse regime, high resolution microscopy based techniques must operate in the ballistic or quasi-ballistic regimes (a few scattering lengths), where the achievable depth is limited by scattering of both the excitation light and the intrinsically generated signal. With laser excitation in the 800–1000 nm range, typical imaging depths in tissues are approximately 100–300 microns. While this range is not highly limiting for imaging ex-vivo physically sliced samples, these depths are inadequate for true in vivo applications. Thus it would be highly advantageous to achieve better imaging depth in nonlinear optical microscopy methods in general. It would be additionally powerful for the specific case of SHG to achieve this goal while simultaneously exploiting the power of polarization analysis to obtain detailed structural data about the tissue assembly.

However performing these measurements in strongly scattering tissues is problematic since each scattering event experienced by both the laser fundamental and SHG wave introduces some depolarization.11 As a result, structural information encoded in SHG will become increasingly scrambled at increasing tissue depths of greater than one mean free path (MFP) for the fundamental and/or SHG signal. While there have been several papers on SHG polarization analysis, these have either been performed at either superficial depths in tissues or in isolated myofibers or collagen fibers.9, 12, 13 The evolution of how these SHG responses degrade in tissues for the excitation and signal waves has not yet been reported. To fully exploit the power of SHG it is important to both determine the usable range as well as increase upon this limit for polarization analysis. As we are limited in terms of both convenient laser sources as well as the biological transparency window in using longer excitation wavelengths, other approaches must be found to increase imaging depths.

To help overcome the inherent depth limitations in turbid tissues while maintaining high resolution coupled with polarization analysis, we will combine SHG imaging with optical clearing.14 Several researchers have recently employed this method in a variety of optical modalities. In this process, a high refractive index, hyper-osmotic reagent (e.g. glycerol, sugars, or sugar alcohols) is added to the tissue to increase its transparency. Such diverse tissues as skin,15-18 blood,19, 20 dura mater,21 gastric tissue,22 sclera,23, 24 bone,25 and muscle26 have been studied by this process, where the optical imaging modalities have included brightfield microscopy, tissue spectroscopy, OCT, and SHG microscopy. In many cases the penetration depth was increased by several fold. It is widely accepted that a decrease in the reduced scattering coefficient, μs’, is responsible for this effect, where the optical clearing potential has been defined as (μsclearedsuncleared).16 As described by Mie theory,27 this ratio becomes infinite for the case of perfect refractive index matching.27

Previously, we investigated the mechanisms and benefits of optical clearing for SHG imaging of tendon and striated muscle.26, 28 We demonstrated that in muscle optical clearing arises from replacement of intracellular water with glycerol (n=1.47) thus matching the initially lower refractive index of the cytoplasm in the muscle cells (n=1.38) with the higher refractive index of the surrounding collagenous perimysium (n=1.47). In the case of tendon in addition to refractive index matching via interfibrillar water replacement with glycerol, this interfibrillar spacing is increased resulting in a longer mean free path (MFP). In these efforts we measured the bulk optical parameters and performed Monte Carlo simulations of the axial directional (Forward/Backward) and attenuation responses and found the experimentally observed improvement in imaging was consistent with the reduction of μs’.28

2. Experimental Methods

2.1. Tissue preparation.

The tendon and muscle samples were obtained from adult CD1 mice sacrificed by CO2 narcosis, in accordance with our approved animal care protocol. To isolate tendon fibrils the skin was pulled from tail, and strips of tendon collagen were carefully detached from bones, and transferred into phosphate buffered saline (PBS). Before imaging tendon was cut into smaller fragments (0.5cm long). Snips of quadriceps femoris muscles were dissected from lower limbs and sliced on a vibratome into 200 μm horizontal sections. Optical clearing was performed overnight in 50% glycerol-PBS solution.

2.2. SHG microscope.

The SHG imaging system consists of a laser scanning unit (Olympus Fluoview 300) mounted on an upright microscope (Olympus BX61), where the excitation source is a mode-locked Titanium Sapphire laser (Coherent Mira). All measurements were taken with a laser fundamental wavelength of 890 nm with average power between 5 and 20 mW at the focal plane. The imaging system simultaneously collects both the forward and the backward components of SHG signal. In the former, a long working distance 40X 0.8 NA water-immersion objective and a 0.9 NA condenser provide excitation and signal collection, respectively. The backward component is collected in a non-descanned configuration. In both geometries, the SHG signal is isolated with a longwave pass dichroic mirror and 10 nm bandpass filters (445 nm). The signals are detected by two identical photon-counting photomultiplier modules (Hamamatsu). The SHG wavelength was confirmed with a fiber optic spectrometer (Ocean Optics). There is no autofluorescence for collagen or skeletal muscle at this excitation wavelength.

The input polarization is controlled by a set of half- and quarter-wave plates, where the latter is used to compensate for ellipticity in the polarization introduced in the beampath. We de facto determined the polarization of excitation light at the focal plane by matching SHG maxima and minima to those previously measured for linear (myofibrils) and spherical (circular cells) specimens. To perform the dependence of the laser polarization at fixed focal planes we simultaneously recorded forward and backward SHG images, while rotating the plane of polarization at 5° steps at high zoom (4x). Changing the polarization by either the half-wave plate or specimen rotation produced similar results. We adapted both acquisition channels for analysis of the polarization of the resulting SHG signals by addition of collimation lenses and Glan laser polarizers. The initial polarization was set to the maximum of the laser polarization response (45 degrees with respect to the fiber axis).9 Images were taken with the analyzing polarizers oriented parallel and orthogonal relative to the laser polarization to calculate the resulting signal anisotropy, β. Using membrane stained spherical neuroblastoma cells we obtained a correction factor of 0.08 for the anisotropy measurement to account for minor losses of polarization introduced by the dichroic mirror in the collection path. For both polarization responses, this analysis is performed for single fibrils or small groups of parallel-oriented fibrils with ImageJ software (http://rsb.info.nih.gov/ij).

3. Results

3.1 Laser polarization dependence of the SHG intensity from tendon

First, we investigate the dependence of the SHG intensity on the laser polarization for uncleared and cleared tendon at different tissue depths. For collagen fibrils, the SHG intensity profile with respect to angle with input polarization, θ, exhibits a angular dependence with maxima at 45 and 135 degrees and local minima at 0, 90, and 180 degrees, respectively.9 The results for control tendon are shown in Fig. 1, where the top panels display representative optical sections for 0, 45, and 90 degrees for 5 and 45 microns into the tissue. The bottom panel plots the intensity dependence on the laser polarization over 180 degrees for these depths. For the 5 um depth (blue squares) the polarization profile is the same as we have previously reported.9 By contrast, the angular dependence is essentially completely lost at a depth of only 45 μm from the tendon surface (red circles). This arises because the laser fundamental gets depolarized by scattering events prior to its arrival at the focal plane, which is equivalent to probing the sample with random polarization. At the laser excitation wavelength of 900 nm, the scattering coefficient μs is 400 cm-1 or equivalently the MFP=25 microns.28 Thus, the laser photons on average will experience two scattering events which results in significant depolarization.

Fig. 1. Angular dependence of the forward SHG intensity from control tendon at depths of 5 and 45 microns. Representative optical sections at several polarization angles are shown in the top panels. Representative error bars (standard error) are given for one laser polarization for each depth.

Fig. 2. Angular dependence of the forward SHG intensity from optically cleared tendon at 10 (black squares) and 100 (red circles) micron depths. Representative optical sections for 100 microns are shown in the top panel Representative error bars (standard error) are given for one laser polarization for each depth.

3.2 Tendon SHG anisotropy

The second polarization response to be examined is the SHG signal anisotropy, which relates to the alignment of dipoles within the focal volume. The polarization state of the SHG signal can be described by its anisotropy parameter β according to Eq. 1:

β=IparIorthIpar+2Iorth
(1)

where Ipar and Iorth are components of SHG intensity polarized parallel and orthogonal with respect to the polarization of excitation laser. The signal anisotropy ranges between -0.5 (all dipoles aligned perpendicular to the laser polarization), and β=1.0 where all the dipoles are aligned with themselves and with the laser polarization. The special case of β=0 corresponds to the isotropic condition where Ipar = Iorth. We will use this scenario to describe the complete depolarization of the SHG signal by scattering events during its propagation through turbid media. Here we will investigate how β evolves with increasing depth (and scattering) for tendon, and further, if it can be retained through optical clearing. This is an important consideration as loss of the SHG anisotropy reduces the information content that SHG provides in comparing differences in tissue structure.

We perform this analysis for both the forward and backward SHG components, as the latter will be the most relevant for in vivo applications. The detected backward SHG is a super-position of direct quasi-coherent emission30 and a multiple scattered incoherent component. In general, the measured signal in turbid tissue results from both contributions,31 where the relative magnitudes depend on the morphology of the tissue assembly as well as the bulk optical parameters. We have determined that the initial emission directionality, FSHG/BSHG, is ~ 5:1 in tendon (un-published results).

To this end, we probed the samples at preset z-positions with fixed vertically polarized laser excitation, and analyzed those tendon fibrils aligned at 45 degrees where the SHG intensity has maximum according to profile in Fig. 1. Each optical section was imaged twice with the Glan polarizers oriented parallel and then perpendicular to the laser polarization. The resulting depth dependence of both the forward and backward collected channels of the anisotropy parameter for tendon over the tendon thickness of 60 microns is shown in Fig. 3.

Fig. 3. SHG anisotropy β measured for forward (red squares) and backward (blue circles) for control tendon.

Fig. 4. (a) Forward SHG anisotropy β measured for forward SHG from control (red squares) and cleared (blue squares) tendon. (b) Forward (red squares) and backward (blue circles) anisotropy in cleared tendon.

3.3 Laser polarization dependence of the SHG intensity from striated muscle

As reported by several researchers, the SHG from striated muscle arises from myosin and has characteristic dependence on the laser polarization, where the angular dependence of the SHG intensity exhibits maxima at 45 and 135 degrees and a minimum at 90 degrees.9, 13 Here we investigate how this response degrades at increasing depths into muscle tissue. Data for the control muscle is shown in Figure 5, along with the cleared data at depths of 10, 100, and 180 microns. We note the input polarization profile is largely randomized at the depth of 100 μm into the muscle tissue. The scattering coefficient at 890 nm for striated muscle is 285 cm-1, thus the MFP is ~35 microns, corresponding to approximately 3 scattering events. This randomizes the laser polarization as we observed in tendon (Figure 1). Upon treatment with 50% glycerol, the polarization profile is preserved through 100 μm of the optically cleared muscle. Some depolarization is evident at 180 μm depth as the amplitude of the curve goes down, especially in the 0 to 45 degrees range where peak-to-valley difference (i.e., dynamic range) is inherently lower. These observations are consistent with the previously measured 10-fold decrease in the scattering coefficient to 29 cm-1 (MFP ~340 microns) in cleared muscle, where no depolarization will occur into 100 microns of tissue thickness (≪MFP) and limited effect in 180 microns (<1 MFP).

Fig. 5. Angular dependence of the forward SHG from control and optically cleared muscle at different depths up to 180 microns.

3.4 SHG anisotropy from striated muscle

In analogy to tendon, we measured the SHG anisotropy with increasing depth for control and optically cleared tendon (Fig. 6). The fibers were oriented at 45 degrees relative to the vertical laser polarization to achieve the greatest intensity. Unlike tendon, muscle tissue did not produce a monotonic depth-dependence of the SHG anisotropy, due to differences in the tissue assembly. Specifically, tendon has uniformly aligned collinear fibrils throughout the whole fiber, where any bending or change in direction occurs gradually with depth. In contrast, the muscle tissue is comprised of multiple cells (~20–50 microns in diameter) wrapped in a thin (~2 μm) layer of collagenous perimysium. The boundary between the muscle cells causes a change in birefringence that will change the polarization of the scattered photons. As discussed by Wang the depolarization in a birefringent medium does not occur isotropically but depends on the initial polarization, the birefringence, and shape of scatterers.32 In the current case, the anisotropy displayed oscillatory behavior at these locations. We previously observed the effects of these local changes in the axial attenuation of the SHG in muscle, where a curvature in the response occurred at the cell boundaries.26 Additionally, artifacts are introduced by slicing at the top and bottom surfaces of the samples, whereas tendon samples were imaged intact. All these factors combined result in abrupt changes in local fibril alignments at and in the vicinity of all the interfaces. Therefore, in Figure 6 we present the average SHG signal anisotropy values from the middle regions of two muscle cells in the biopsy at average depths of 50 and 180 microns. Similar to tendon, the uncleared tissue shows significant depolarization at the top and bottom of the tissue. There is a slight increase in the anisotropy at the bottom exit as the laser depolarization of the laser is not as strong as observed in tendon (based on the respective scattering coefficients at 890 nm).

Fig. 6. SHG anisotropy β calculated for forward SHG from control (red squares) and cleared (blue circles) muscle samples. The data is presented at two depths corresponding to the middle regions of two muscle cells at 50 and 180 microns.

4. Discussion

We recognize that there still might be potential safety issues with the use of clearing agents in vivo both in terms of toxicity and exposure time. As described above, both tendon and muscle swell upon treatment with 50% glycerol. Other researchers have reported shrinking under glycerol treatment due to de-hydration in the inter-celullar space.35 The difference in behavior is likely due to the cellularity of the tissue structure. For example, tendon is completely a-cellular and skeletal muscle consists of tightly packed supercells wrapped with collagen and essentially no inter-cellular space for hydration. In contrast, gastric tissue has highly dynamic water flow. We further note that development of optical clearing methods36 is an active area of research and new agents or delivery schemes may be found that are suitable for in vivo use. However, the ability to gain further knowledge into tissue structure may provide insight into changes that occur during connective tissue disorders and cancer. Moreover, imaging ex vivo tissues through optical clearing offers advantages over standard histology. Sections for histology require significant preparation in terms of fixing, imbedding in paraffin, and then slicing, where by contrast through SHG imaging and optical clearing, much thicker sections (~200 microns) can be used. As a consequence we can obtain much more data from one section than possible by ~20 thin histological sections. Thus optical clearing coupled with SHG polarization analysis is a powerful method even if it is ultimately limited to ex vivo applications.

5. Conclusions

In this work we investigated the effect of optical clearing on polarization properties of SHG emission from highly scattering muscle and tendon samples. To this end, we measured the dependence of SHG intensity on the laser polarization angle and SHG signal anisotropy at different depths. We find that in untreated tissues, based on our measured bulk optical parameters, the polarization signatures are highly randomized in 2-3 scattering events. In cleared tissues, we found that the SHG polarization information is retained in ~200 μm-thick samples, where these effects are consistent with the reduction of the scattering coefficients (and corresponding mean free paths). These findings have significance for probing tissue structure through polarization analysis at increased depths relative to the superficial depths possible of un-treated tissues. While we demonstrated retention of polarization attributes through optical clearing for the specific case of SHG imaging, this method could be applied to other polarization sensitive contrast modalities like fluorescence third harmonic generation, coherent anti-Stokes Raman scattering, and optical coherence tomography.

Acknowledgments

We gratefully acknowledge support under NIH EB01842 and helpful discussions with Dr. Ron LaComb.

References and links

1.

O. Nadiarnykh, S. Plotnikov, W. A. Mohler, I. Kalajzic, D. Redford-Badwal, and P. J. Campagnola, “Second Harmonic Generation imaging microscopy studies of Osteogenesis Imperfecta,” J. Biomed. Opt. 12, 051805 (2007). [CrossRef] [PubMed]

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4.

E. Brown, T. McKee, E. diTomaso, A. Pluen, B. Seed, Y. Boucher, and R. K. Jain, “Dynamic imaging of collagen and its modulation in tumors in vivo using second-harmonic generation,” Nat. Med. 9(6), 796–800 (2003). [CrossRef] [PubMed]

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P. P. Provenzano, K. W. Eliceiri, J. M. Campbell, D. R. Inman, J. G. White, and P. J. Keely, “Collagen reorganization at the tumor-stromal interface facilitates local invasion,” BMC Med. 4, 38 (2006). [CrossRef] [PubMed]

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M. C. Skala, K. M. Riching, D. K. Bird, A. Gendron-Fitzpatrick, J. Eickhoff, K. W. Eliceiri, P. J. Keely, and N. Ramanujam, “In vivo multiphoton fluorescence lifetime imaging of protein-bound and free nicotinamide adenine dinucleotide in normal and precancerous epithelia,” J. Biomed. Opt. 12, 024014 (2007). [CrossRef] [PubMed]

9.

S. V. Plotnikov, A. C. Millard, P. J. Campagnola, and W. A. Mohler, “Characterization of the Myosin-based source for second-harmonic generation from muscle sarcomeres,” Biophys J. 90, 693–703 (2006). [CrossRef]

10.

P. J. Campagnola, A. C. Millard, M. Terasaki, P. E. Hoppe, C. J. Malone, and W. A. Mohler, “3-Dimesional High-Resolution Second Harmonic Generation Imaging of Endogenous Structural Proteins in Biological Tissues,” Biophys. J. 82, 493–508 (2002). [CrossRef]

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M. Moscoso, J. B. Keller, and G. Papanicolaou, “Depolarization and blurring of optical images by biological tissue,” J. Opt. Soc. Am. A 18, 948–960 (2001). [CrossRef]

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C.-W. Sun, C.-Y. Wang, C. C. Yang, Y.-W. Kiang, I-. Hsu, and C.-W. Lin, “Polarization gating in ultrafast-optics imaging of skeletal muscle tissues,” Opt. Lett. 26, 432–434 (2001). [CrossRef]

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S.-W. Chu, S.-Y. Chen, G.-W. Chern, T.-H. Tsai, Y.-C. Chen, B.-L. Lin, and C.-K. Sun, “Studies of (2)/(3) Tensors in Submicron-Scaled Bio-Tissues by Polarization Harmonics Optical Microscopy,” Biophys. J. 86, 3914–3922 (2004). [CrossRef] [PubMed]

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V. Tuchin, Optical Clearing of Tissues and Blood (SPIE Press, Bellingham, WA, 2006), Vol. PM 154.

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X. Q. Xu, R. K. Wang, and J. B. Elder, “Optical clearing effect on gastric tissues immersed with biocompatible chemical agents investigated by near infrared reflectance spectroscopy,” J. Phys. D 36, 1707–1713 (2003). [CrossRef]

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M. V. Schulmerich, J. H. Cole, K. A. Dooley, M. D. Morris, J. M. Kreider, and S. A. Goldstein, “Optical clearing in transcutaneous Raman spectroscopy of murine cortical bone tissue,” J. Biomed. Opt. 13, 021108 (2008). [CrossRef] [PubMed]

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28.

R. LaComb, O. Nadiarnykh, S. Carey, and P. J. Campagnola, “Quantitative SHG imaging and modeling of the optical clearing mechanism in striated muscle and tendon,” J. Biomed. Opt. 13, 021109 (2008). [CrossRef] [PubMed]

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A. T. Yeh and J. Hirshburg, “Molecular interactions of exogenous chemical agents with collagen--implications for tissue optical clearing,” J Biomed. Opt. 11, 014003 (2006). [CrossRef] [PubMed]

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R. Lacomb, O. Nadiarnykh, S. S. Townsend, and P. J. Campagnola, “Phase Matching considerations in Second Harmonic Generation from tissues: Effects on emission directionality, conversion efficiency and observed morphology,” Opt. Comm. 281, 1823–1832 (2008). [CrossRef]

31.

O. Nadiarnykh, R. B. LaComb, P. J. Campagnola, and W. A. Mohler, “Coherent and incoherent SHG in fibrillar cellulose matrices,” Opt. Express 15, 3348–3360 (2007). [CrossRef] [PubMed]

32.

X. Wang and L. V. Wang, “Propagation of polarized light in birefringent turbid media: a Monte Carlo study,” J. Biomed. Opt. 7, 279–290 (2002). [CrossRef] [PubMed]

33.

P. Stoller, B.-M. Kim, A. M. Rubinchik, K. M. Reiser, and L. B. Da Silva, “Polarization-dependent optical second-harmonic imaging of a rat-tail tendon,” J. Biomed. Opt. 7, 205–214 (2001). [CrossRef]

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C. E. Bigelow and T. H. Foster, “Confocal fluorescence polarization microscopy in turbid media: effects of scattering-induced depolarization,” J. Opt. Soc. Am. A 23, 2932–2943 (2006). [CrossRef]

35.

R. K. K. Wang, X. Q. Xu, Y. H. He, and J. B. Elder, “Investigation of optical clearing of gastric tissue immersed with hyperosmotic agents,” IEEE J. Select. Top. Quantum Electron. 9, 234–242 (2003). [CrossRef]

36.

J. Yoon, T. Son, E. H. Choi, B. Choi, J. S. Nelson, and B. Jung, “Enhancement of optical skin clearing efficacy using a microneedle roller,” J. Biomed. Opt. 13, 021103 (2008). [CrossRef] [PubMed]

OCIS Codes
(180.6900) Microscopy : Three-dimensional microscopy
(190.2620) Nonlinear optics : Harmonic generation and mixing
(190.4180) Nonlinear optics : Multiphoton processes
(290.4210) Scattering : Multiple scattering
(290.7050) Scattering : Turbid media

ToC Category:
Microscopy

History
Original Manuscript: December 15, 2008
Revised Manuscript: January 16, 2009
Manuscript Accepted: January 19, 2009
Published: March 26, 2009

Virtual Issues
Vol. 4, Iss. 5 Virtual Journal for Biomedical Optics

Citation
Oleg Nadiarnykh and Paul J. Campagnola, "Retention of polarization signatures in SHG microscopy of scattering tissues through optical clearing," Opt. Express 17, 5794-5806 (2009)
http://www.opticsinfobase.org/oe/abstract.cfm?URI=oe-17-7-5794


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References

  1. O. Nadiarnykh, S. Plotnikov, W. A. Mohler, I. Kalajzic, D. Redford-Badwal, and P. J. Campagnola, "Second Harmonic Generation imaging microscopy studies of Osteogenesis Imperfecta," J. Biomed. Opt. 12, 051805 (2007). [CrossRef] [PubMed]
  2. R. Lacomb, O. Nadiarnykh, and P. J. Campagnola, "Quantitative SHG imaging of the diseased state Osteogenesis Imperfecta: Experiment and Simulation," Biophys J 94, 4104- (2008). [CrossRef]
  3. S. V. Plotnikov, A. Kenny, S. Walsh, B. Zubrowski, C. Joseph, V. L. Scranton, G. A. Kuchel, D. Dauser, M. Xu, C. Pilbeam, D. Adams, R. Dougherty, P. J. Campagnola, and W. A. Mohler, "Measurement of muscle disease by quantitative second-harmonic generation imaging," J. Biomed. Opt. 13, 044018 (2008). [CrossRef] [PubMed]
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