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Optics Express

Optics Express

  • Editor: C. Martijn de Sterke
  • Vol. 19, Iss. 8 — Apr. 11, 2011
  • pp: 7587–7595
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Single-shot Full-field reflection phase microscopy

Zahid Yaqoob, Toyohiko Yamauchi, Wonshik Choi, Dan Fu, Ramachandra R. Dasari, and Michael S. Feld  »View Author Affiliations


Optics Express, Vol. 19, Issue 8, pp. 7587-7595 (2011)
http://dx.doi.org/10.1364/OE.19.007587


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Abstract

We present a full-field reflection phase microscope that combines low-coherence interferometry and off-axis digital holographic microscopy (DHM). The reflection-based DHM provides highly sensitive and a single-shot imaging of cellular dynamics while the use of low coherence source provides a depth-selective measurement. The setup uniquely uses a diffraction grating in the reference arm to generate an interference image of uniform contrast over the entire field-of-view albeit low-coherence light source. We have measured the path-length sensitivity of our instrument to be approximately 21 p i c o m e t e r s / H z that makes it suitable for nanometer-scale full-field measurement of membrane dynamics in live cells.

© 2011 OSA

1. Introduction

Bio-microrheology is the quantitative study of mechanical properties of live cells [1

1. D. Weihs, T. G. Mason, and M. A. Teitell, “Bio-microrheology: a frontier in microrheology,” Biophys. J. 91(11), 4296–4305 (2006). [CrossRef] [PubMed]

]. Variations in mechanical properties are intrinsic indicators of ongoing cellular processes such as increase in elasticity of certain cancer cells [2

2. M. Beil, A. Micoulet, G. von Wichert, S. Paschke, P. Walther, M. B. Omary, P. P. Van Veldhoven, U. Gern, E. Wolff-Hieber, J. Eggermann, J. Waltenberger, G. Adler, J. Spatz, and T. Seufferlein, “Sphingosylphosphorylcholine regulates keratin network architecture and visco-elastic properties of human cancer cells,” Nat. Cell Biol. 5(9), 803–811 (2003). [CrossRef] [PubMed]

], change of membrane stiffness in malaria-infected red blood cells [3

3. Y. K. Park, M. Diez-Silva, G. Popescu, G. Lykotrafitis, W. S. Choi, M. S. Feld, and S. Suresh, “Refractive index maps and membrane dynamics of human red blood cells parasitized by Plasmodium falciparum,” Proc. Natl. Acad. Sci. U.S.A. 105(37), 13730–13735 (2008). [CrossRef] [PubMed]

], changes in cellular adhesion [4

4. N. Almqvist, R. Bhatia, G. Primbs, N. Desai, S. Banerjee, and R. Lal, “Elasticity and adhesion force mapping reveals real-time clustering of growth factor receptors and associated changes in local cellular rheological properties,” Biophys. J. 86(3), 1753–1762 (2004). [CrossRef] [PubMed]

], and so forth. The measurement of rheological properties of cell membranes is advantageous since it may also indirectly provide information on the internal structures of cell [1

1. D. Weihs, T. G. Mason, and M. A. Teitell, “Bio-microrheology: a frontier in microrheology,” Biophys. J. 91(11), 4296–4305 (2006). [CrossRef] [PubMed]

]. A number of different techniques exist to assess membrane rheological properties of live cells. These include atomic force microscopy (AFM) [5

5. J. Alcaraz, L. Buscemi, M. Grabulosa, X. Trepat, B. Fabry, R. Farre, and D. Navajas, “Microrheology of human lung epithelial cells measured by atomic force microscopy,” Biophys. J. 84(3), 2071–2079 (2003). [CrossRef] [PubMed]

], optical and magnetic tweezers [6

6. M. Puig-de-Morales-Marinkovic, K. T. Turner, J. P. Butler, J. J. Fredberg, and S. Suresh, “Viscoelasticity of the human red blood cell,” Am. J. Physiol. Cell Physiol. 293(2), C597–C605 (2007). [CrossRef] [PubMed]

, 7

7. K. Svoboda and S. M. Block, “Biological applications of optical forces,” Annu. Rev. Biophys. Biomol. Struct. 23(1), 247–285 (1994). [CrossRef] [PubMed]

], pipette aspiration [8

8. E. Evans and A. Leung, “Adhesivity and rigidity of erythrocyte membrane in relation to wheat germ agglutinin binding,” J. Cell Biol. 98(4), 1201–1208 (1984). [CrossRef] [PubMed]

10

10. K. G. Engstrom, B. Moller, and H. J. Meiselman, “Optical Evaluation of Red Blood Cell Geometry Using Micropipette aspiration,” Blood Cells 18(2), 241–257, discussion 258–265 (1992). [PubMed]

], electric field deformation [11

11. H. Engelhardt, H. Gaub, and E. Sackmann, “Viscoelastic properties of Erythrocyte Membranes in High-Frequency Electric Fields,” Nature 307(5949), 378–380 (1984). [CrossRef] [PubMed]

], and full-field transmission phase microscopy [12

12. G. Popescu, T. Ikeda, R. R. Dasari, and M. S. Feld, “Diffraction phase microscopy for quantifying cell structure and dynamics,” Opt. Lett. 31(6), 775–777 (2006). [CrossRef] [PubMed]

]. Many of these methods are invasive and use large deformations that may lead to cell damage or cell’s response to mechanical strain rather than its intrinsic response. For point-measurement techniques such as AFM, the time scales to probe large surface areas of cell membrane are in minutes, preventing the study of high-speed cell membrane dynamics over a wider surface area. Full-field transmission phase microscopy has been successfully utilized to measure membrane flcutuations in red blood cells over a broad range of temporal and spatial frequencies [12

12. G. Popescu, T. Ikeda, R. R. Dasari, and M. S. Feld, “Diffraction phase microscopy for quantifying cell structure and dynamics,” Opt. Lett. 31(6), 775–777 (2006). [CrossRef] [PubMed]

]. Furthermore, the measured membrane fluctuations can be used in appropriate mathematical models to calculate rheological properties of red blood cells [13

13. Y. Park, C. A. Best, K. Badizadegan, R. R. Dasari, M. S. Feld, T. Kuriabova, M. L. Henle, A. J. Levine, and G. Popescu, “Measurement of red blood cell mechanics during morphological changes,” Proc. Natl. Acad. Sci. U.S.A. 107(15), 6731–6736 (2010). [CrossRef] [PubMed]

, 14

14. R. Wang, H. Ding, M. Mir, K. Tangella, and G. Popescu, “Effective 3D viscoelasticity of red blood cells measured by diffraction phase microscopy,” Biomed. Opt. Express 2(3), 485–490 (2011). [CrossRef] [PubMed]

]. However, for most type of cells, which have complicated 3-D internal cellular structures, transmission-type optical techniques will not be suitable as they will probe a combination of membrane as well as bulk properties of cells that are difficult to decouple. In this context, properly designed reflection-based phase microscopy with depth-sectioning capability can play vital role to exclusively access the membrane dynamics of nucleated cells. Moreover, transmission mode techniques measure relative phase shift induced by the sample with respect to that by the medium. Thus, the measured phase shift is proportional to the refractive index difference, Δn, between the sample and the medium. In contrast, reflection phase microscopy techniques yield phase measurement proportional to the index of refraction, n, of the medium rather than the relative index, Δn. Thus, reflection-based optical methods promise a 2nn advantage in measurement sensitivity over the transmission-based optical techniques.

Low-coherence interferometry is necessary to exclusively sample the reflection signal from the depth of interest. In the past, both spectral domain as well as time domain optical coherence tomography (OCT) based implementations of reflection phase microscopy have been reported [15

15. M. A. Choma, A. K. Ellerbee, C. Yang, T. L. Creazzo, and J. A. Izatt, “Spectral-domain phase microscopy,” Opt. Lett. 30(10), 1162–1164 (2005). [CrossRef] [PubMed]

19

19. Z. Yaqoob, W. Choi, S. Oh, N. Lue, Y. Park, C. Fang-Yen, R. R. Dasari, K. Badizadegan, and M. S. Feld, “Improved phase sensitivity in spectral domain phase microscopy using line-field illumination and self phase-referencing,” Opt. Express 17(13), 10681–10687 (2009). [CrossRef] [PubMed]

]. Joo et al. and Choma et al. have independently developed similar setups using self phase-referenced spectral domain phase microscopy setups using point illumination [15

15. M. A. Choma, A. K. Ellerbee, C. Yang, T. L. Creazzo, and J. A. Izatt, “Spectral-domain phase microscopy,” Opt. Lett. 30(10), 1162–1164 (2005). [CrossRef] [PubMed]

, 16

16. C. Joo, T. Akkin, B. Cense, B. H. Park, and J. F. de Boer, “Spectral-domain optical coherence phase microscopy for quantitative phase-contrast imaging,” Opt. Lett. 30(16), 2131–2133 (2005). [CrossRef] [PubMed]

]. Ellerbee et al. used the phase sensitive OCT based configuration in [15

15. M. A. Choma, A. K. Ellerbee, C. Yang, T. L. Creazzo, and J. A. Izatt, “Spectral-domain phase microscopy,” Opt. Lett. 30(10), 1162–1164 (2005). [CrossRef] [PubMed]

] to visualize the motion of intracellular structures [20

20. A. K. Ellerbee, T. L. Creazzo, and J. A. Izatt, “Investigating nanoscale cellular dynamics with cross-sectional spectral domain phase microscopy,” Opt. Express 15(13), 8115–8124 (2007). [CrossRef] [PubMed]

]. In recent past, we have designed and developed a quantitative phase microscope based on spectral domain OCT and line-field illumination [19

19. Z. Yaqoob, W. Choi, S. Oh, N. Lue, Y. Park, C. Fang-Yen, R. R. Dasari, K. Badizadegan, and M. S. Feld, “Improved phase sensitivity in spectral domain phase microscopy using line-field illumination and self phase-referencing,” Opt. Express 17(13), 10681–10687 (2009). [CrossRef] [PubMed]

]. The line-field reflection phase microscope exploited low-coherent illumination and confocal gating to successfully obtain the surface profile of cell membrane with sub-nanometer axial resolution. Using the line-field approach, we demonstrated 1 kHz frame rate with more than hundred data points along the line illumination. The first full-field phase sensitive OCT was reported using swept-source OCT configuration, which required 1024 wavelength encoded images to generate a volume phase image [17

17. M. V. Sarunic, S. Weinberg, and J. A. Izatt, “Full-field swept-source phase microscopy,” Opt. Lett. 31(10), 1462–1464 (2006). [CrossRef] [PubMed]

]. Moreover, the acquisition rate (25 ms integration time per wavelength) was not sufficient to observe cellular dynamics. In order to observe intrinsic membrane motion of living cells, Yamauchi et al. developed a full-field time-domain reflection phase microscope based on phase shifting interferometry and captured sectional surface profile of living cells. But the time resolution was limited to 1.25 sec due to the need for taking multiple images [18

18. T. Yamauchi, H. Iwai, M. Miwa, and Y. Yamashita, “Low-coherent quantitative phase microscope for nanometer-scale measurement of living cells morphology,” Opt. Express 16(16), 12227–12238 (2008). [CrossRef] [PubMed]

]. There was an attempt to use an off-axis digital holography with low-coherence source to take a full-field phase image in a single shot [21

21. P. Massatsch, F. Charriere, E. Cuche, P. Marquet, and C. D. Depeursinge, “Time-domain optical coherence tomography with digital holographic microscopy,” Appl. Opt. 44(10), 1806–1812 (2005). [CrossRef] [PubMed]

]. But the tilting of reference mirror caused uneven interference contrast and thereby impeded full-field imaging.

2. Full-field reflection phase microscope

2.1 Experimental setup

Figure 1
Fig. 1 (a) Schematic of full-field single-shot reflection phase microscope. SMF: single mode fiber, Li: ith spherical lens, BSi: ith beam splitter, G: diffraction grating, Si: ith spatial filter. (b) Typical interferogram with a flat surface as the sample after subtracting the no fringe image representing the DC signal.
shows the schematic of our single-shot full-field reflection phase microscope (FF-RPM). Light from a mode-locked Ti:Sapphire laser (center wavelength, λc = 800 nm) is coupled into a single-mode fiber for delivery as well as for spectrum broadening. The full-width-half-maximum spectral width, Δλ, at the fiber output measures 50 nm, which yields a round trip coherence length of 4 μm in a typical culture medium with refractive index, n, equal to 1.33. The sample beam that travels through lenses L2, L3, L4, and a water immersion objective lens L5 (Olympus UPlanSApo 60×/1.2 W), reflects off the sample surface and makes an image of the sample on a high-speed complementary metal oxide semiconductor (CMOS) camera (Photron 1024PCI) via L6 and L15. The reference beam, which passes through L7, L8, L9 and L10, is diverted on its way back using a beam splitter BS2. Portion of the reference beam that goes back through BS2 is blocked using a spatial filter S1. On the other hand, the deflected beam passes through L11-L14 and combines with the returning sample beam at the 3rd beam splitter BS3. For off-axis interferometry, a diffraction grating G is introduced in one of the conjugate planes. Out of multiple diffracted orders, we choose only the +1st order by placing a spatial filter S2 in the Fourier plane of lens L12. As a result, the diffracted reference beam interferes with the sample beam in the image plane at an angle. We note that the period of the diffraction grating and the magnification between the grating and the camera provide the desired angular shift to the reference beam for off-axis interferometry. Since the grating and the camera suffice the imaging condition, the optical path length measured from any point on the grating to the corresponding pixel on the camera is constant. As a result, we achieve homogeneous fringe visibility across the whole field-of-view unlike the setup in [21

21. P. Massatsch, F. Charriere, E. Cuche, P. Marquet, and C. D. Depeursinge, “Time-domain optical coherence tomography with digital holographic microscopy,” Appl. Opt. 44(10), 1806–1812 (2005). [CrossRef] [PubMed]

] that simply used reference mirror tilt for off-axis interferometry. We note that the proposed setup is capable of taking quantitative phase images in double-pass transmission mode [22

22. H. Iwai, C. Fang-Yen, G. Popescu, A. Wax, K. Badizadegan, R. R. Dasari, and M. S. Feld, “Quantitative phase imaging using actively stabilized phase-shifting low-coherence interferometry,” Opt. Lett. 29(20), 2399–2401 (2004). [CrossRef] [PubMed]

] as well as reflection mode, which is achieved by placing the coherence gate on the glass slide or the cell membrane, respectively.

2.2 Signal processing

Figure 1(b) shows a measured interferogram with a flat surface as the sample. As expected, the spatial fringes are straight as well as equally spaced when the sample is flat. The total measured intensity at the CMOS camera can be written as
I(x,y)=IR+IS(x,y)+2IRIS(x,y)cos[ux+vy+φ(x,y)],
(1)
where IR and IS(x,y)are the reference and sample beam intensity distributions, respectively. u and v represent the frequency of spatial fringes along the x- and y- axes, and φ(x,y) is the spatially varying phase associated with the sample under study. We also acquire a no-fringe image that represents the DC component in Eq. (1) by shifting the coherence gate out of the sample. By subtracting the no-fringe image from the original interferogram, we get only the interference term.

3. Results and discussion

3.1 Common mode noise rejection and measurement sensitivity

Intrinsic membrane fluctuations in living cells are typically on the order of a nanometer or less; the measurement of these small membrane fluctuations requires the development of quantitative phase microscopes with high signal-to-noise ratio (SNR). In this section, we show the measurement sensitivity of our full-field RPM in terms of the least detectable axial motion; the configuration to measure the measurement sensitivity is shown in Fig. 3(a)
Fig. 3 (a) Configuration to determine the sensitivity of FF-RPM. and (b) Measured phase fluctuation (radian) as a function of applied voltage. Mi: ith mirror, PZT: Lead Zirconate Titanate.
. The full-field illumination shines on both the surfaces; mirror M1 mounted on a translation stage and mirror M2 attached to a Lead Zirconate Titanate (PZT) actuator.

In order to suppress the common mode noise due to independent mechanical or thermal fluctuations of the reference beam path with respect to the sample beam path, we utilize a self-phase referencing method which is previously described in Ref [19

19. Z. Yaqoob, W. Choi, S. Oh, N. Lue, Y. Park, C. Fang-Yen, R. R. Dasari, K. Badizadegan, and M. S. Feld, “Improved phase sensitivity in spectral domain phase microscopy using line-field illumination and self phase-referencing,” Opt. Express 17(13), 10681–10687 (2009). [CrossRef] [PubMed]

]. Since the phase of all the points in the full-field illumination is acquired at the same time, every point in the field of view shares the same interferometric noise as any other point. We take the phase measured from a portion of the beam illuminating the reflector M1 as the reference phase, representing the common-mode noise. By subtracting this reference phase from the phase of the subsequent points on M2, we remove the common-mode noise to obtain actual fluctuation of the surface M2.

3.2 Quantitative phase imaging of live cells

The advantage of the reflection-mode imaging is also evident when comparing Eqs. (3) and (4) in the context of SNR. In other words, assuming that the phase sensitivity of the transmission and reflection-mode measurements is same, the height resolution (or measurement sensitivity) of the reflection phase imaging is 40 times (nm/Δn) better than that of transmission measurement. Moreover, the reflection phase image will reveal the shape of the cell surface independent of the distribution of intracellular refractive index since it depends only on the refractive index of the medium which can be accurately measured by a conventional refractometer.

3.3 Measurement of cell membrane fluctuation

As discussed in section 1, membrane fluctuations are intrinsic indicator of overall cellular condition and have been used in the past to successfully estimate membrane mechanical properties in relation to different stages of malaria infection in human red blood cells [3

3. Y. K. Park, M. Diez-Silva, G. Popescu, G. Lykotrafitis, W. S. Choi, M. S. Feld, and S. Suresh, “Refractive index maps and membrane dynamics of human red blood cells parasitized by Plasmodium falciparum,” Proc. Natl. Acad. Sci. U.S.A. 105(37), 13730–13735 (2008). [CrossRef] [PubMed]

]. But for eukaryotic cells having complex internal structures, our full-field reflection phase microscope is well-suited to selectively measure membrane fluctuations by effectively choosing to reject contributions from the internal cellular structures.

To demonstrate our system’s capabilities, we have studied the membrane fluctuations in HeLa cells under different cell conditions. More specifically, we prepared (i) a sample of living normal HeLa cells, (ii) a fixed HeLa cell sample after treatment with 2% paraformaldehyde and (iii) a sample of HeLa cells treated with 8 nM Cytochalasin-D which inhibits actin polymerization [25

25. J. F. Casella, M. D. Flanagan, and S. Lin, “Cytochalasin D inhibits actin polymerization and induces depolymerization of actin filaments formed during platelet shape change,” Nature 293(5830), 302–305 (1981). [CrossRef] [PubMed]

]. The frame rate of the image acquisition was set to 1 kHz and the data was recorded for duration of 1 sec for each cell. As shown in Fig. 5(a)
Fig. 5 Setup and results of the cell membrane fluctuation measurement. (a) Location of coherence gate; the sample is tilted to simultaneously acquire membrane fluctuations as well as background phase from the coverslip. (b) Power spectral density of membrane fluctuations as a function of frequency for three different populations: blue, formalin fixed; green, normal; and red, Cytochalasin-D treated HeLa cells.
, the sample under test was tilted to simultaneously acquire membrane fluctuations as well as background phase from the coverslip. By subtracting the background phase change observed on the coverslip, the common-mode mechanical noise was eliminated. We measured the temporal fluctuations on the cell surface and calculated the PSD of membrane motion for each cell. Figure 5(b) shows the mean PSD for each cell population. The number of normal, fixed, and Cytochalasin-D treated cells used in this study were N = 22, 20, and 33, respectively. As expected, the PSD of the fixed cells was measured smaller and flatter than the normal ones implying that the cell membrane became stiffer after chemical fixation. On the other hand, the PSD of the Cytochalasin-D treated cells was measured larger than the normal ones implying that the cell membrane became softer due to the inhibition of actin polymerization.

4. Conclusion

We have proposed and demonstrated, for the first time, a quantitative reflection phase microscope based on en-face optical coherence tomography and off-axis digital holography. The setup utilizes a diffraction grating in the reference arm to provide the desired angular tilt to the reference beam for off-axis interferometry. The full-field illumination allows single-shot phase measurement of multiple points on the surface of interest and enables the use of self phase-referencing method to reject common-mode noise inherent in interferometric setups using a separate reference arm. In our full-field reflection phase microscope, the self-phase referencing suppressed phase detection noise down to as low as 21picometers/Hz. With such high phase sensitivity, we were able to resolve thermal motion of the cell surface in the field of view, which was on the order of 100 picometers to 150 nanometers. A potential application of the full-field reflection phase microscope is to use the membrane fluctuations to estimate the mechanical properties of cell membrane; these variations in cell membrane mechanical properties can serve as non-invasive biomarker to study pathophysiology of general cell types [13

13. Y. Park, C. A. Best, K. Badizadegan, R. R. Dasari, M. S. Feld, T. Kuriabova, M. L. Henle, A. J. Levine, and G. Popescu, “Measurement of red blood cell mechanics during morphological changes,” Proc. Natl. Acad. Sci. U.S.A. 107(15), 6731–6736 (2010). [CrossRef] [PubMed]

]. Another future direction includes full-field and multi-cell imaging of cellular electromotility, including cell membrane motion driven by the action potential in single mammalian cells [26

26. P. C. Zhang, A. M. Keleshian, and F. Sachs, “Voltage-induced membrane movement,” Nature 413(6854), 428–432 (2001). [CrossRef] [PubMed]

].

Acknowledgements

This work was funded by the National Center for Research Resources of the National Institutes of Health (P41-RR02594-25), the National Science Foundation (DBI-0754339), and Hamamatsu Photonics.

References and links

1.

D. Weihs, T. G. Mason, and M. A. Teitell, “Bio-microrheology: a frontier in microrheology,” Biophys. J. 91(11), 4296–4305 (2006). [CrossRef] [PubMed]

2.

M. Beil, A. Micoulet, G. von Wichert, S. Paschke, P. Walther, M. B. Omary, P. P. Van Veldhoven, U. Gern, E. Wolff-Hieber, J. Eggermann, J. Waltenberger, G. Adler, J. Spatz, and T. Seufferlein, “Sphingosylphosphorylcholine regulates keratin network architecture and visco-elastic properties of human cancer cells,” Nat. Cell Biol. 5(9), 803–811 (2003). [CrossRef] [PubMed]

3.

Y. K. Park, M. Diez-Silva, G. Popescu, G. Lykotrafitis, W. S. Choi, M. S. Feld, and S. Suresh, “Refractive index maps and membrane dynamics of human red blood cells parasitized by Plasmodium falciparum,” Proc. Natl. Acad. Sci. U.S.A. 105(37), 13730–13735 (2008). [CrossRef] [PubMed]

4.

N. Almqvist, R. Bhatia, G. Primbs, N. Desai, S. Banerjee, and R. Lal, “Elasticity and adhesion force mapping reveals real-time clustering of growth factor receptors and associated changes in local cellular rheological properties,” Biophys. J. 86(3), 1753–1762 (2004). [CrossRef] [PubMed]

5.

J. Alcaraz, L. Buscemi, M. Grabulosa, X. Trepat, B. Fabry, R. Farre, and D. Navajas, “Microrheology of human lung epithelial cells measured by atomic force microscopy,” Biophys. J. 84(3), 2071–2079 (2003). [CrossRef] [PubMed]

6.

M. Puig-de-Morales-Marinkovic, K. T. Turner, J. P. Butler, J. J. Fredberg, and S. Suresh, “Viscoelasticity of the human red blood cell,” Am. J. Physiol. Cell Physiol. 293(2), C597–C605 (2007). [CrossRef] [PubMed]

7.

K. Svoboda and S. M. Block, “Biological applications of optical forces,” Annu. Rev. Biophys. Biomol. Struct. 23(1), 247–285 (1994). [CrossRef] [PubMed]

8.

E. Evans and A. Leung, “Adhesivity and rigidity of erythrocyte membrane in relation to wheat germ agglutinin binding,” J. Cell Biol. 98(4), 1201–1208 (1984). [CrossRef] [PubMed]

9.

R. P. Hebbel, A. Leung, and N. Mohandas, “Oxidation-induced changes in microrheologic properties of the red blood cell membrane,” Blood 76(5), 1015–1020 (1990). [PubMed]

10.

K. G. Engstrom, B. Moller, and H. J. Meiselman, “Optical Evaluation of Red Blood Cell Geometry Using Micropipette aspiration,” Blood Cells 18(2), 241–257, discussion 258–265 (1992). [PubMed]

11.

H. Engelhardt, H. Gaub, and E. Sackmann, “Viscoelastic properties of Erythrocyte Membranes in High-Frequency Electric Fields,” Nature 307(5949), 378–380 (1984). [CrossRef] [PubMed]

12.

G. Popescu, T. Ikeda, R. R. Dasari, and M. S. Feld, “Diffraction phase microscopy for quantifying cell structure and dynamics,” Opt. Lett. 31(6), 775–777 (2006). [CrossRef] [PubMed]

13.

Y. Park, C. A. Best, K. Badizadegan, R. R. Dasari, M. S. Feld, T. Kuriabova, M. L. Henle, A. J. Levine, and G. Popescu, “Measurement of red blood cell mechanics during morphological changes,” Proc. Natl. Acad. Sci. U.S.A. 107(15), 6731–6736 (2010). [CrossRef] [PubMed]

14.

R. Wang, H. Ding, M. Mir, K. Tangella, and G. Popescu, “Effective 3D viscoelasticity of red blood cells measured by diffraction phase microscopy,” Biomed. Opt. Express 2(3), 485–490 (2011). [CrossRef] [PubMed]

15.

M. A. Choma, A. K. Ellerbee, C. Yang, T. L. Creazzo, and J. A. Izatt, “Spectral-domain phase microscopy,” Opt. Lett. 30(10), 1162–1164 (2005). [CrossRef] [PubMed]

16.

C. Joo, T. Akkin, B. Cense, B. H. Park, and J. F. de Boer, “Spectral-domain optical coherence phase microscopy for quantitative phase-contrast imaging,” Opt. Lett. 30(16), 2131–2133 (2005). [CrossRef] [PubMed]

17.

M. V. Sarunic, S. Weinberg, and J. A. Izatt, “Full-field swept-source phase microscopy,” Opt. Lett. 31(10), 1462–1464 (2006). [CrossRef] [PubMed]

18.

T. Yamauchi, H. Iwai, M. Miwa, and Y. Yamashita, “Low-coherent quantitative phase microscope for nanometer-scale measurement of living cells morphology,” Opt. Express 16(16), 12227–12238 (2008). [CrossRef] [PubMed]

19.

Z. Yaqoob, W. Choi, S. Oh, N. Lue, Y. Park, C. Fang-Yen, R. R. Dasari, K. Badizadegan, and M. S. Feld, “Improved phase sensitivity in spectral domain phase microscopy using line-field illumination and self phase-referencing,” Opt. Express 17(13), 10681–10687 (2009). [CrossRef] [PubMed]

20.

A. K. Ellerbee, T. L. Creazzo, and J. A. Izatt, “Investigating nanoscale cellular dynamics with cross-sectional spectral domain phase microscopy,” Opt. Express 15(13), 8115–8124 (2007). [CrossRef] [PubMed]

21.

P. Massatsch, F. Charriere, E. Cuche, P. Marquet, and C. D. Depeursinge, “Time-domain optical coherence tomography with digital holographic microscopy,” Appl. Opt. 44(10), 1806–1812 (2005). [CrossRef] [PubMed]

22.

H. Iwai, C. Fang-Yen, G. Popescu, A. Wax, K. Badizadegan, R. R. Dasari, and M. S. Feld, “Quantitative phase imaging using actively stabilized phase-shifting low-coherence interferometry,” Opt. Lett. 29(20), 2399–2401 (2004). [CrossRef] [PubMed]

23.

T. Ikeda, G. Popescu, R. R. Dasari, and M. S. Feld, “Hilbert phase microscopy for investigating fast dynamics in transparent systems,” Opt. Lett. 30(10), 1165–1167 (2005). [CrossRef] [PubMed]

24.

W. Choi, C. Fang-Yen, K. Badizadegan, S. Oh, N. Lue, R. R. Dasari, and M. S. Feld, “Tomographic phase microscopy,” Nat. Methods 4(9), 717–719 (2007). [CrossRef] [PubMed]

25.

J. F. Casella, M. D. Flanagan, and S. Lin, “Cytochalasin D inhibits actin polymerization and induces depolymerization of actin filaments formed during platelet shape change,” Nature 293(5830), 302–305 (1981). [CrossRef] [PubMed]

26.

P. C. Zhang, A. M. Keleshian, and F. Sachs, “Voltage-induced membrane movement,” Nature 413(6854), 428–432 (2001). [CrossRef] [PubMed]

OCIS Codes
(120.3890) Instrumentation, measurement, and metrology : Medical optics instrumentation
(170.1530) Medical optics and biotechnology : Cell analysis
(170.4500) Medical optics and biotechnology : Optical coherence tomography

ToC Category:
Medical Optics and Biotechnology

History
Original Manuscript: February 7, 2011
Revised Manuscript: March 17, 2011
Manuscript Accepted: March 18, 2011
Published: April 5, 2011

Virtual Issues
Vol. 6, Iss. 5 Virtual Journal for Biomedical Optics

Citation
Zahid Yaqoob, Toyohiko Yamauchi, Wonshik Choi, Dan Fu, Ramachandra R. Dasari, and Michael S. Feld, "Single-shot Full-field reflection phase microscopy," Opt. Express 19, 7587-7595 (2011)
http://www.opticsinfobase.org/oe/abstract.cfm?URI=oe-19-8-7587


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References

  1. D. Weihs, T. G. Mason, and M. A. Teitell, “Bio-microrheology: a frontier in microrheology,” Biophys. J. 91(11), 4296–4305 (2006). [CrossRef] [PubMed]
  2. M. Beil, A. Micoulet, G. von Wichert, S. Paschke, P. Walther, M. B. Omary, P. P. Van Veldhoven, U. Gern, E. Wolff-Hieber, J. Eggermann, J. Waltenberger, G. Adler, J. Spatz, and T. Seufferlein, “Sphingosylphosphorylcholine regulates keratin network architecture and visco-elastic properties of human cancer cells,” Nat. Cell Biol. 5(9), 803–811 (2003). [CrossRef] [PubMed]
  3. Y. K. Park, M. Diez-Silva, G. Popescu, G. Lykotrafitis, W. S. Choi, M. S. Feld, and S. Suresh, “Refractive index maps and membrane dynamics of human red blood cells parasitized by Plasmodium falciparum,” Proc. Natl. Acad. Sci. U.S.A. 105(37), 13730–13735 (2008). [CrossRef] [PubMed]
  4. N. Almqvist, R. Bhatia, G. Primbs, N. Desai, S. Banerjee, and R. Lal, “Elasticity and adhesion force mapping reveals real-time clustering of growth factor receptors and associated changes in local cellular rheological properties,” Biophys. J. 86(3), 1753–1762 (2004). [CrossRef] [PubMed]
  5. J. Alcaraz, L. Buscemi, M. Grabulosa, X. Trepat, B. Fabry, R. Farre, and D. Navajas, “Microrheology of human lung epithelial cells measured by atomic force microscopy,” Biophys. J. 84(3), 2071–2079 (2003). [CrossRef] [PubMed]
  6. M. Puig-de-Morales-Marinkovic, K. T. Turner, J. P. Butler, J. J. Fredberg, and S. Suresh, “Viscoelasticity of the human red blood cell,” Am. J. Physiol. Cell Physiol. 293(2), C597–C605 (2007). [CrossRef] [PubMed]
  7. K. Svoboda and S. M. Block, “Biological applications of optical forces,” Annu. Rev. Biophys. Biomol. Struct. 23(1), 247–285 (1994). [CrossRef] [PubMed]
  8. E. Evans and A. Leung, “Adhesivity and rigidity of erythrocyte membrane in relation to wheat germ agglutinin binding,” J. Cell Biol. 98(4), 1201–1208 (1984). [CrossRef] [PubMed]
  9. R. P. Hebbel, A. Leung, and N. Mohandas, “Oxidation-induced changes in microrheologic properties of the red blood cell membrane,” Blood 76(5), 1015–1020 (1990). [PubMed]
  10. K. G. Engstrom, B. Moller, and H. J. Meiselman, “Optical Evaluation of Red Blood Cell Geometry Using Micropipette aspiration,” Blood Cells 18(2), 241–257, discussion 258–265 (1992). [PubMed]
  11. H. Engelhardt, H. Gaub, and E. Sackmann, “Viscoelastic properties of Erythrocyte Membranes in High-Frequency Electric Fields,” Nature 307(5949), 378–380 (1984). [CrossRef] [PubMed]
  12. G. Popescu, T. Ikeda, R. R. Dasari, and M. S. Feld, “Diffraction phase microscopy for quantifying cell structure and dynamics,” Opt. Lett. 31(6), 775–777 (2006). [CrossRef] [PubMed]
  13. Y. Park, C. A. Best, K. Badizadegan, R. R. Dasari, M. S. Feld, T. Kuriabova, M. L. Henle, A. J. Levine, and G. Popescu, “Measurement of red blood cell mechanics during morphological changes,” Proc. Natl. Acad. Sci. U.S.A. 107(15), 6731–6736 (2010). [CrossRef] [PubMed]
  14. R. Wang, H. Ding, M. Mir, K. Tangella, and G. Popescu, “Effective 3D viscoelasticity of red blood cells measured by diffraction phase microscopy,” Biomed. Opt. Express 2(3), 485–490 (2011). [CrossRef] [PubMed]
  15. M. A. Choma, A. K. Ellerbee, C. Yang, T. L. Creazzo, and J. A. Izatt, “Spectral-domain phase microscopy,” Opt. Lett. 30(10), 1162–1164 (2005). [CrossRef] [PubMed]
  16. C. Joo, T. Akkin, B. Cense, B. H. Park, and J. F. de Boer, “Spectral-domain optical coherence phase microscopy for quantitative phase-contrast imaging,” Opt. Lett. 30(16), 2131–2133 (2005). [CrossRef] [PubMed]
  17. M. V. Sarunic, S. Weinberg, and J. A. Izatt, “Full-field swept-source phase microscopy,” Opt. Lett. 31(10), 1462–1464 (2006). [CrossRef] [PubMed]
  18. T. Yamauchi, H. Iwai, M. Miwa, and Y. Yamashita, “Low-coherent quantitative phase microscope for nanometer-scale measurement of living cells morphology,” Opt. Express 16(16), 12227–12238 (2008). [CrossRef] [PubMed]
  19. Z. Yaqoob, W. Choi, S. Oh, N. Lue, Y. Park, C. Fang-Yen, R. R. Dasari, K. Badizadegan, and M. S. Feld, “Improved phase sensitivity in spectral domain phase microscopy using line-field illumination and self phase-referencing,” Opt. Express 17(13), 10681–10687 (2009). [CrossRef] [PubMed]
  20. A. K. Ellerbee, T. L. Creazzo, and J. A. Izatt, “Investigating nanoscale cellular dynamics with cross-sectional spectral domain phase microscopy,” Opt. Express 15(13), 8115–8124 (2007). [CrossRef] [PubMed]
  21. P. Massatsch, F. Charriere, E. Cuche, P. Marquet, and C. D. Depeursinge, “Time-domain optical coherence tomography with digital holographic microscopy,” Appl. Opt. 44(10), 1806–1812 (2005). [CrossRef] [PubMed]
  22. H. Iwai, C. Fang-Yen, G. Popescu, A. Wax, K. Badizadegan, R. R. Dasari, and M. S. Feld, “Quantitative phase imaging using actively stabilized phase-shifting low-coherence interferometry,” Opt. Lett. 29(20), 2399–2401 (2004). [CrossRef] [PubMed]
  23. T. Ikeda, G. Popescu, R. R. Dasari, and M. S. Feld, “Hilbert phase microscopy for investigating fast dynamics in transparent systems,” Opt. Lett. 30(10), 1165–1167 (2005). [CrossRef] [PubMed]
  24. W. Choi, C. Fang-Yen, K. Badizadegan, S. Oh, N. Lue, R. R. Dasari, and M. S. Feld, “Tomographic phase microscopy,” Nat. Methods 4(9), 717–719 (2007). [CrossRef] [PubMed]
  25. J. F. Casella, M. D. Flanagan, and S. Lin, “Cytochalasin D inhibits actin polymerization and induces depolymerization of actin filaments formed during platelet shape change,” Nature 293(5830), 302–305 (1981). [CrossRef] [PubMed]
  26. P. C. Zhang, A. M. Keleshian, and F. Sachs, “Voltage-induced membrane movement,” Nature 413(6854), 428–432 (2001). [CrossRef] [PubMed]

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