## The wide-field optical sectioning of microlens array and structured illumination-based plane-projection multiphoton microscopy |

Optics Express, Vol. 21, Issue 2, pp. 2097-2109 (2013)

http://dx.doi.org/10.1364/OE.21.002097

Acrobat PDF (2302 KB)

### Abstract

We present a theoretical investigation of an optical microscope design that achieves wide-field, multiphoton fluorescence microscopy with finer axial resolution than confocal microscopy. Our technique creates a thin plane of excitation light at the sample using height-staggered microlens arrays (HSMAs), wherein the height staggering of microlenses generate temporal focusing to suppress out-of-focus excitation, and the dense spacing of microlenses enables the implementation of structured illumination technique to eliminate residual out-of-focus signal. We use physical optics-based numerical simulations to demonstrate that our proposed technique can achieve diffraction-limited three-dimensional imaging through a simple optical design.

© 2013 OSA

## 1. Introduction

1. J. B. Pawley, *Handbook of Biological Confocal Microscopy*, 3rd ed. (Springer, 2006). [CrossRef]

4. D. Oron, E. Tal, and Y. Silberberg, “Scanningless depth-resolved microscopy,” Opt. Express **13**, 1468–1476 (2005). [CrossRef] [PubMed]

2. W. Denk, J. Strickler, and W. Webb, “2-photon laser scanning fluorescence microscopy,” Science **248**, 73–76 (1990). [CrossRef] [PubMed]

3. J. Huisken, J. Swoger, F. Del Bene, J. Wittbrodt, and E. Stelzer, “Optical sectioning deep inside live embryos by selective plane illumination microscopy,” Science **305**, 1007–1009 (2004). [CrossRef] [PubMed]

6. M. Neil, R. Juskaitis, and T. Wilson, “Method of obtaining optical sectioning by using structured light in a conventional microscope,” Opt. Lett. **22**, 1905–1907 (1997). [CrossRef]

1. J. B. Pawley, *Handbook of Biological Confocal Microscopy*, 3rd ed. (Springer, 2006). [CrossRef]

7. B. Masters, P. So, C. Buehler, N. Barry, J. Sutin, W. Mantulin, and E. Gratton, “Mitigating thermal mechanical damage potential during two-photon dermal imaging,” J. Biomed. Opt. **9**, 1265–1270 (2004). [CrossRef] [PubMed]

9. J. Bewersdorf, R. Pick, and S. Hell, “Multifocal multiphoton microscopy,” Opt. Lett. **23**, 655–657 (1998). [CrossRef]

10. V. Andresen, A. Egner, and S. Hell, “Time-multiplexed multifocal multiphoton microscope,” Opt. Lett. **26**, 75–77 (2001). [CrossRef]

^{2}-focus area, see Appendix A, B), which limits the degree of parallelization. SIM and SPIM do not suffer from parallelization issues; however both techniques have their own limitations. SIM can lead to significant photobleaching and low signal-to-noise ratio (SNR) in the processed images, because it excites fluorophores and receives fluorescence over a wide axial range at each acquisition. SPIM requires a compromise between axial resolution and the size of the field of view. Moreover, SPIM raises design issues and challenges in sample handling and manipulation for its separate excitation and imaging objectives in close proximity.

11. P. J. Keller, A. D. Schmidt, A. Santella, K. Khairy, Z. Bao, J. Wittbrodt, and E. H. K. Stelzer, “Fast, high-contrast imaging of animal development with scanned light sheet-based structured-illumination microscopy,” Nat. Methods **7**, 637–U55 (2010). [CrossRef] [PubMed]

4. D. Oron, E. Tal, and Y. Silberberg, “Scanningless depth-resolved microscopy,” Opt. Express **13**, 1468–1476 (2005). [CrossRef] [PubMed]

## 2. MASI-PPMP: integrating (intrinsic) temporal focusing and (extrinsic) SIM

12. J.-Y. Yu, C.-H. Kuo, D. B. Holland, Y. Chen, M. Ouyang, G. A. Blake, R. Zadoyan, and C.-L. Guo, “Wide-field optical sectioning for live-tissue imaging by plane-projection multiphoton microscopy,” J. Biomed. Opt. **16**, 116009 (2011). [CrossRef] [PubMed]

13. C. Ventalon and J. Mertz, “Quasi-confocal fluorescence sectioning with dynamic speckle illumination,” Opt. Lett. **30**, 3350–3352 (2005). [CrossRef]

6. M. Neil, R. Juskaitis, and T. Wilson, “Method of obtaining optical sectioning by using structured light in a conventional microscope,” Opt. Lett. **22**, 1905–1907 (1997). [CrossRef]

14. A. Egner and S. Hell, “Time multiplexing and parallelization in multifocal multiphoton microscopy,” J. Opt. Soc. Am. A **17**, 1192–1201 (2000). [CrossRef]

*t*in TM-MMM and MASI-PPMP are different. In TM-MMM, Δ

*t*is set to be much larger than the pulse duration

*τ*

_{0}to reduce temporal inter-foci interferences [14

14. A. Egner and S. Hell, “Time multiplexing and parallelization in multifocal multiphoton microscopy,” J. Opt. Soc. Am. A **17**, 1192–1201 (2000). [CrossRef]

*t*<

*τ*

_{0}for practical reasons detailed below and in Supplementary Materials.

*d*is the aperture diameter of a single time-delay channel,

*λ*

_{0}is the central wavelength of the excitation light (here ∼ 800 nm), and Δ

*h*is the height staggering between the longest and shortest time-delay channels. Accordingly, we restrict Δ

_{max}*h*to ∼ 300

_{max}*μ*m and 36

*μ*m ≥

*d*≥ 18

*μ*m, via Eq. (1). These dimensions make the HSMA fabrication feasible through existing techniques, and can create up to ∼ 10

^{6}foci within a 2-inch aperture, thereby providing the periodic, high-spatial-frequency pattern for SIM. For a simple analysis, we assume that the total amount of time delay Δ

*t*is separated equally into

_{tot}*N*distinct time-delay steps (with step size Δ

_{t}*t*, Eq. (2)). These time-delay steps are then arranged in a prescribed periodic pattern in the HSMA (Fig. 1(b)). Considering the propagation speed of light in a material, we have where

*c*and

*n*are the speed of light in vacuum and the refractive index of the material of the HSMAs (set to be 1.5), respectively.

*h*, the estimated Δ

_{max}*t*in Eq. (2) can be around or shorter than the pulse duration of conventional ultrafast oscillators or amplifiers above certain values of

*N*. In such cases, we should take into account the temporal interferences among light pulses of different time delays. Notably, temporal masks with much larger Δ

_{t}*t*have been proposed to avoid temporal interferences and to achieve scanningless TM-MMM [14

_{tot}14. A. Egner and S. Hell, “Time multiplexing and parallelization in multifocal multiphoton microscopy,” J. Opt. Soc. Am. A **17**, 1192–1201 (2000). [CrossRef]

*h*falls far beyond the limits of existing fabrication techniques, and the aperture sizes of these temporal masks could be too large for standard biomedical microscopes (see Appendix B).

_{max}*N*(and Δ

_{t}*t*) for a given Δ

*t*(§4), and the arrangement of time delays in the HSMA (Appendix C). Then, we couple HSMAs with SIM to enhance optical sectioning (§5).

_{tot}## 3. Construct a physical optics-based model taking into account temporal interferences

**17**, 1192–1201 (2000). [CrossRef]

*E*(

**r**,

*z*,

*t*) at position (

**r**,

*z*) (

*z*= 0 at the specimen plane) and time

*t*is approximated as the Gaussian-weighted sum of a series of constant-interval (in

*k*–space), in-phase light waves, where

*k*

_{0}is the central wavenumber of the pulse spectrum, and

*E*

_{kj}is the scalar field of the light wave of wavenumber

*k*. To approximate the ultrafast pulse train generated by the amplified laser system we used previously [12

_{j}12. J.-Y. Yu, C.-H. Kuo, D. B. Holland, Y. Chen, M. Ouyang, G. A. Blake, R. Zadoyan, and C.-L. Guo, “Wide-field optical sectioning for live-tissue imaging by plane-projection multiphoton microscopy,” J. Biomed. Opt. **16**, 116009 (2011). [CrossRef] [PubMed]

*k*

_{0}≈ 7.85 × 10

^{4}cm

^{−1}and a pulse duration

*τ*

_{0}of ≈ 30 fs (by using an appropriate

*σ*). We then employ the amplitude point spread function (PSF) derived previously for high numerical-aperture (NA) lenses [14

_{k}**17**, 1192–1201 (2000). [CrossRef]

*E*

_{kj}, as where

*α*is the maximal focusing angle

*θ*of the objective lens, and

*J*

_{0}is the zero-order Bessel function of the first kind. The objective lens used in all the simulations presented here is a 60X oil-immersion lens of NA 1.42.

*E*(

_{PSF}**r**,

*z*,

*t*). Having solved

*E*numerically, we estimate the electric field near the specimen plane,

_{PSF}*E*(

_{SP}**r**,

*z*,

*t*), as the linear superposition of the

*E*from the individual microlenses, where

_{PSF}**r**and Δ

_{m}*t*

**are the central position and time delay of the ultrafast pulse going though the**

_{m}**m**-th HSMA channel, respectively. An example of numerically simulated

*E*(

_{SP}**r**,

*z*,

*t*) is shown in Media 1, which also reveals the dynamic process of temporal focusing (see Appendix D). Notably, to fulfill the wide-field illumination condition and to simplify the simulations, our model assumes that a ’unit’ microlens array is infinitely replicated in the transverse coordinates, as shown in Fig. 1(b). Under such a periodic condition, the physical optics properties in the projected region of one unit microlens array is sufficient to represent the entire system (the validity and details of this model are given in Appendix E).

*I*(

_{SP}**r**,

*z*) can be derived by integrating the excitation intensity over time, as where

*n*is the number of photons required in single excitation event (here

_{p}*n*= 2).

_{p}## 4. Optimize intrinsic optical sectioning through tuning *N*_{t} and Δ*t*

_{t}

*z*,

*S*(

*z*), through integration of

*I*(

_{SP}**r**,

*z*) in Eq. (6) over the transverse coordinates, as Experimentally,

*S*(

*z*) corresponds to the detected fluorescence signal from a thin fluorescent film placed at a depth

*z*.

*N*, Δ

_{t}*t*) that produces the most efficient optical sectioning, we evaluate

*S*(

*z*) and the ratio of out-of-focus and in-focus signal,

*S*/

_{out}*S*, for various sets of

_{in}*N*and Δ

_{t}*t*(constrained by Eq. (2)) with two inter-foci spacings,

*d*(≡

_{foci}*d*/

*M*, where

*M*is the magnification of the microscopy system) ≈ 0.4 and 0.8

*λ*

_{0}, corresponding to ∼ 56 and ∼ 14 foci per 10

^{2}-focus area, respectively (Fig. 2). Because of the square geometry of our HSMAs, we examine

*N*= 2

_{t}^{2}, 3

^{2}, 4

^{2}, ... and 9

^{2}. Analyzing

*S*/

_{out}*S*reveals that the decay of the out-of-focus excitation significantly slows down between

_{in}*N*= 25 and 64, corresponding to Δ

_{t}*t*≈ 20-8 fs (i.e., 2/3

*τ*

_{0}-1/4

*τ*

_{0}) (Fig. 2(c)). We observe similar trends of

*S*/

_{out}*S*for various geometrical arrangements of the distinct time-delay steps (see Appendix C). This result suggests that the intrinsic optical sectioning for a fixed Δ

_{in}*t*is optimized when Δ

_{tot}*t*is slightly smaller than

*τ*

_{0}. Further reducing Δ

*t*(equivalent to increasing

*N*(Eq. (2))) can complicate the fabrication of the HSMAs without major improvement of the intrinsic optical sectioning. In addition, increasing

_{t}*d*leads to weaker out-of-focus excitation and a less complex axial excitation profile

_{foci}*S*(

*z*) (Fig. 2(b)).

## 5. Obtain extrinsic optical sectioning via implementing SIM

*I*. Because the emitted fluorescence (wavelength assumed to be ∼ 0.56

_{IM}*λ*

_{0}) from the specimen is generally incoherent, we can estimate

*I*from a convolution of the excitation intensity profile

_{IM}*I*and the intensity PSF of the microscopy system,

_{SP}*I*[5], as where

_{SYS}*f*is the concentration distribution of the fluorophore in the specimen. To quantify the optical-sectioning effect, we assume that the specimen is an ideal thin fluorescent film placed at

*z*=

*z*, i.e.,

_{f}*f*(r,

*z*) =

*δ*(

*z*−

*z*). Eq. (8) then becomes Conventional SIM takes 3 shifts of the 1-dimension periodic pattern; each step is 1/3 of the period of the pattern [6

_{f}6. M. Neil, R. Juskaitis, and T. Wilson, “Method of obtaining optical sectioning by using structured light in a conventional microscope,” Opt. Lett. **22**, 1905–1907 (1997). [CrossRef]

*I*is 2-dimensional. We thus use 3-by-3 shifts (

_{SP}*I*

_{IM}_{1},

*I*

_{IM}_{2}, ...,

*I*

_{IM}_{9}denote the obtained images) and apply SIM post-processing to extract the optically sectioned images, We then substitute

*I*in Eq. (7) with

_{SP}*I*to evaluate the overall strength of optical sectioning (Fig. 3).

_{SIM}*I*. Other methods using a high-spatial-frequency illumination pattern to distinguish in-focus and out-of-focus signals, such as HiLo microscopy [17

_{IM}17. D. Lim, K. K. Chu, and J. Mertz, “Wide-field fluorescence sectioning with hybrid speckle and uniform-illumination microscopy,” Opt. Lett. **33**, 1819–21 (2008). [CrossRef] [PubMed]

18. D. Lim, T. N. Ford, K. K. Chu, and J. Mertz, “Optically sectioned in vivo imaging with speckle illumination hilo microscopy,” J. Biomed. Opt. **16**, 016014 (2011). [CrossRef] [PubMed]

## 6. Conclusion

^{2}-focus area), and thus has greater potential in high-speed imaging. In addition to the finer axial resolution than CFM, the lateral resolution can be enhanced via utilization of the periodic excitation structures and Fourier analysis of the obtained images [19

19. R. Heintzmann and P. A. Benedetti, “High-resolution image reconstruction in fluorescence microscopy with patterned excitation,” Appl. Opt. **45**, 5037–5045 (2006). [CrossRef] [PubMed]

12. J.-Y. Yu, C.-H. Kuo, D. B. Holland, Y. Chen, M. Ouyang, G. A. Blake, R. Zadoyan, and C.-L. Guo, “Wide-field optical sectioning for live-tissue imaging by plane-projection multiphoton microscopy,” J. Biomed. Opt. **16**, 116009 (2011). [CrossRef] [PubMed]

## Appendix A: Determining the dimensions of the time-delay channels

*h*, as shown in Fig. 5. When the light of wavelength

*λ*

_{0}exits from channel 1 (Ch1) and propagates to distance

*z*, diffraction theory predicts the beam size,

*d*, as a function of

_{B}*λ*

_{0}and

*z*. With this regard, we consider the inter-channel leakage to be non-negligible if

*d*before propagating to

_{B}*z*= Δ

*h*is larger than the diameter of the microlens,

*d*. In other words, the inter-channel leakage is negligible if A reasonable design, as shown in Fig. 5, is to set the focus of Ch1 at

*z*= Δ

*h*, so that the

*d*decreases upon exiting Ch1 and reaches its minimum,

_{B}*d*, at

_{f}*z*= Δ

*h*. Based on the diffraction-limited spot size of an ideal thin lens, we can estimate

*d*as where

_{f}*f*

_{#}is defined as

*f*/

*d*[20]. Combining Eqs. (11), (12) and the geometry, we have and the criterion of negligible inter-channel light leakage is thus To ensure the inter-channel light leakage is negligible regardless of the geometrical arrangements of distinct time-delay steps, we can substitute the largest height difference in the HSMA, Δ

*h*, for Δ

_{max}*h*in Eq. (14), and thus derive

**17**, 1192–1201 (2000). [CrossRef]

## Appendix B: Size of the microlens arrays to achieve scanningless imaging using conventional time-multiplexing approach

*d*

_{0}, which is commonly much larger than the size of the focal spot of the objective lens,

*d*. As a result, there will be substantial un-illuminated area between the foci. For example, take an oil-immersion, NA 1.42 objective lens: the resultant

_{fs}*d*

_{0}and

*d*is approximately 5

_{fs}*λ*[14

**17**, 1192–1201 (2000). [CrossRef]

*λ*, which makes the fraction of un-illuminated area approximately

^{2}-focus area. To cover the un-illuminated area, Egner et al. [14

**17**, 1192–1201 (2000). [CrossRef]

*N*, can be estimated as

_{t}*N*, and using the propagation speed of light in the HSMA material, we can estimate the difference of height between the longest and shortest time channels, Δ

_{t}*h*, as: where Δ

_{max}*t*,

*c*and

*n*are the step size of the distinct time-delay steps, speed of light in vacuum and the refractive index of the material of the HSMAs, respectively. The appropriate values of Δ

*t*for negligible temporal interferences, as suggested in the previous study of TM-MMM [14

**17**, 1192–1201 (2000). [CrossRef]

*τ*

_{0}, i.e., Δ

*t*≥ 2

*τ*

_{0}(where

*τ*

_{0}was set as ≈ 100 fs for conventional ultrafast oscillators) [14

**17**, 1192–1201 (2000). [CrossRef]

*h*≈ 33 mm and

*d*≈ 0.15 mm using Eqs. (15)–(17), wherein

*λ*and

*n*are assumed to be ∼ 800 nm and 1.5, respectively. As a result, if one needs 1000-by-1000 foci in the field of view (FOV), the typical aperture of the entire HSMA will be as large as 1000×

*d*≈ 150 mm, which is considerably larger than the optical elements of a standard biomedical microscope. Furthermore, when an ultrafast pulse propagates in a long, narrow glass cylinder, both group velocity dispersion and modal dispersion broaden the pulse width, and thus reduce the axial resolution as well as excitation efficiency. From Eqs. (15), (17) and the requirement of Δ

*t*≥ 2

*τ*

_{0}, we can derive that Δ

*h*∝

*τ*

_{0}and

*h*and

*d*can be scaled down via using small

*τ*

_{0}. However, commonly available ultrafast systems typically achieve

*τ*

_{0}≥ 10 fs, such that the resultant Δ

*h*and

*d*remain considerably large. Moreover, the ultrafast pulses of smaller

*τ*

_{0}(and thus greater spectral bandwidth) will suffer more from group velocity dispersion and modal dispersion while propagating in the HSMA materials.

## Appendix C: Various geometrical arrangements of distinct time-delay steps show a similar trend of *S*_{out} / *S*_{in}

_{out}

_{in}

*S*/

_{out}*S*, using various geometrical arrangements of distinct time-delay steps. As concluded in the main article, these

_{in}*S*/

_{out}*S*curves show that the decays of the out-of-focus excitation slow down significantly when

_{in}*N*= 25–64, (Δ

_{t}*t*≈ 2/3

*τ*

_{0}-1/4

*τ*

_{0}), although the curves from different patterns of time delays show slight differences in the values of

*S*/

_{out}*S*.

_{in}## Appendix D: Simulation of the dynamic *E*_{SP} in space reveals temporal focusing

_{SP}

*E*. In particular, we plot the distributions of the real part of the electric field (Re(

_{SP}*E*), upper panel) and intensity (|

_{SP}*E*|

_{SP}^{2}, lower panel). The geometrical arrangement of the distinct time-delay steps used for this simulation is a spiral pattern (Fig. 1(b) of main article) with

*N*= 49 and

_{t}*d*≈ 0.4

_{foci}*λ*

_{0}. Media 1 shows that at most of the area, except for the in-focus region (

*z*≈ 0), the intensity is moderate during nearly the entire time course (100–700 fs). On the other hand, at in-focus region the intensity is high in a relatively short period of time (450–540 fs). Such a phenomenon is commonly recognized as temporal focusing [4

4. D. Oron, E. Tal, and Y. Silberberg, “Scanningless depth-resolved microscopy,” Opt. Express **13**, 1468–1476 (2005). [CrossRef] [PubMed]

## Appendix E: Validity of the assumption of infinitely replicated unit HSMA in a wide-field microscope

*I*at arbitrary positions, the contributions from very distant foci are negligible. In other words, one can approximate the results of using the infinitely replicated unit HSMAs via consideration of only the contributions from the foci within a certain range. To justify such an approach, we need to examine if the computed

_{SP}*I*becomes stationary (i.e., converging to a certain value) as the range of foci considered to contribute, defined by

_{SP}*r*(Fig. 8(a)), is expanded.

_{tot}*I*using Eq. (6) of the main article, with only the contributions to

*E*from the foci within a distance

_{SP}*r*considered (Fig. 8(a)). Here we use

_{tot}*I*, the intensity at a given position as derived with a relatively large

_{end}*r*(≈ 375

_{tot}*λ*

_{0}, where

*λ*

_{0}is the central wavelength of the light pulse), as a reference value, and plot

*I*/

*I*as

_{end}*r*increases from 0 to 125

_{tot}*λ*

_{0}. Fig. 8(b) shows that values of

*I*at all of the 100 randomly chosen positions converge to their corresponding

*I*as

_{end}*r*increases. In particular, we note that when

_{tot}*r*≥ 62.5

_{tot}*λ*

_{0}(equivalent to 50

*μ*m in physical dimensions), the error of

*I*, defined as |

*I*−

*I*|/

_{end}*I*, is less than 0.1%. Such results suggest that, for a given error tolerance in numerical simulations, we can use the inf-HSMA model by considering only the contributions of foci within a certain distance

_{end}*r*. Here, we have

_{inf}*r*≈ 50

_{inf}*μ*m for a 0.1% error tolerance. In the physical microscopy system, we can also use

*r*to determine the region wherein the inf-HSMA assumption is valid, as shown in Fig. 8(c). For conventional biomedical microscopes using

_{inf}*M*= 60X objective lenses, the diameter of the full FOV is typically lager than 300

*μ*m. Thus, the inf-HSMA model is valid in the central region of diameter larger than 200

*μ*m. At the image plane, this region corresponds to a disk of diameter ∼ 12 mm (200

*μ*m×

*M*) or larger, which is able to cover most conventional imaging devices.

## Acknowledgment

## References and links

1. | J. B. Pawley, |

2. | W. Denk, J. Strickler, and W. Webb, “2-photon laser scanning fluorescence microscopy,” Science |

3. | J. Huisken, J. Swoger, F. Del Bene, J. Wittbrodt, and E. Stelzer, “Optical sectioning deep inside live embryos by selective plane illumination microscopy,” Science |

4. | D. Oron, E. Tal, and Y. Silberberg, “Scanningless depth-resolved microscopy,” Opt. Express |

5. | T. Wilson, |

6. | M. Neil, R. Juskaitis, and T. Wilson, “Method of obtaining optical sectioning by using structured light in a conventional microscope,” Opt. Lett. |

7. | B. Masters, P. So, C. Buehler, N. Barry, J. Sutin, W. Mantulin, and E. Gratton, “Mitigating thermal mechanical damage potential during two-photon dermal imaging,” J. Biomed. Opt. |

8. | I. Akira, T. Takeo, I. Katsumi, S. Yumiko, K. Yasuhito, M. Kenta, A. Michio, and U. Isao, “High-speed confocal fluorescence microscopy using a nipkow scanner with microlenses for 3-d imaging of single fluorescent molecule in real time,” Bioimaging |

9. | J. Bewersdorf, R. Pick, and S. Hell, “Multifocal multiphoton microscopy,” Opt. Lett. |

10. | V. Andresen, A. Egner, and S. Hell, “Time-multiplexed multifocal multiphoton microscope,” Opt. Lett. |

11. | P. J. Keller, A. D. Schmidt, A. Santella, K. Khairy, Z. Bao, J. Wittbrodt, and E. H. K. Stelzer, “Fast, high-contrast imaging of animal development with scanned light sheet-based structured-illumination microscopy,” Nat. Methods |

12. | J.-Y. Yu, C.-H. Kuo, D. B. Holland, Y. Chen, M. Ouyang, G. A. Blake, R. Zadoyan, and C.-L. Guo, “Wide-field optical sectioning for live-tissue imaging by plane-projection multiphoton microscopy,” J. Biomed. Opt. |

13. | C. Ventalon and J. Mertz, “Quasi-confocal fluorescence sectioning with dynamic speckle illumination,” Opt. Lett. |

14. | A. Egner and S. Hell, “Time multiplexing and parallelization in multifocal multiphoton microscopy,” J. Opt. Soc. Am. A |

15. | J. Jahns and K.-H. Brenner, |

16. | M. Born and E. Wolf, |

17. | D. Lim, K. K. Chu, and J. Mertz, “Wide-field fluorescence sectioning with hybrid speckle and uniform-illumination microscopy,” Opt. Lett. |

18. | D. Lim, T. N. Ford, K. K. Chu, and J. Mertz, “Optically sectioned in vivo imaging with speckle illumination hilo microscopy,” J. Biomed. Opt. |

19. | R. Heintzmann and P. A. Benedetti, “High-resolution image reconstruction in fluorescence microscopy with patterned excitation,” Appl. Opt. |

20. | E. Hecht, |

**OCIS Codes**

(170.0110) Medical optics and biotechnology : Imaging systems

(180.6900) Microscopy : Three-dimensional microscopy

(260.1960) Physical optics : Diffraction theory

(180.4315) Microscopy : Nonlinear microscopy

**ToC Category:**

Microscopy

**History**

Original Manuscript: October 31, 2012

Revised Manuscript: December 23, 2012

Manuscript Accepted: December 26, 2012

Published: January 18, 2013

**Virtual Issues**

Vol. 8, Iss. 2 *Virtual Journal for Biomedical Optics*

**Citation**

Jiun-Yann Yu, Daniel B. Holland, Geoffrey A. Blake, and Chin-Lin Guo, "The wide-field optical sectioning of microlens array and structured illumination-based plane-projection multiphoton microscopy," Opt. Express **21**, 2097-2109 (2013)

http://www.opticsinfobase.org/oe/abstract.cfm?URI=oe-21-2-2097

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### References

- J. B. Pawley, Handbook of Biological Confocal Microscopy, 3rd ed. (Springer, 2006). [CrossRef]
- W. Denk, J. Strickler, and W. Webb, “2-photon laser scanning fluorescence microscopy,” Science248, 73–76 (1990). [CrossRef] [PubMed]
- J. Huisken, J. Swoger, F. Del Bene, J. Wittbrodt, and E. Stelzer, “Optical sectioning deep inside live embryos by selective plane illumination microscopy,” Science305, 1007–1009 (2004). [CrossRef] [PubMed]
- D. Oron, E. Tal, and Y. Silberberg, “Scanningless depth-resolved microscopy,” Opt. Express13, 1468–1476 (2005). [CrossRef] [PubMed]
- T. Wilson, Confocal Microscopy (Academic Press, 1990).
- M. Neil, R. Juskaitis, and T. Wilson, “Method of obtaining optical sectioning by using structured light in a conventional microscope,” Opt. Lett.22, 1905–1907 (1997). [CrossRef]
- B. Masters, P. So, C. Buehler, N. Barry, J. Sutin, W. Mantulin, and E. Gratton, “Mitigating thermal mechanical damage potential during two-photon dermal imaging,” J. Biomed. Opt.9, 1265–1270 (2004). [CrossRef] [PubMed]
- I. Akira, T. Takeo, I. Katsumi, S. Yumiko, K. Yasuhito, M. Kenta, A. Michio, and U. Isao, “High-speed confocal fluorescence microscopy using a nipkow scanner with microlenses for 3-d imaging of single fluorescent molecule in real time,” Bioimaging4, 57–62 (1996-06).
- J. Bewersdorf, R. Pick, and S. Hell, “Multifocal multiphoton microscopy,” Opt. Lett.23, 655–657 (1998). [CrossRef]
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