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Abstract
Optically shifting the focal plane to allow depth scanning of delicate biological structures and processes in their natural environment offers an appealing alternative to conventional mechanical scanning. Our technique uses a deformable mirror-based photoacoustic remote sensing microscopy (PARS) with a focus shifting of Δz ∼ 240 µm. We achieve this by integrating a deformable mirror that functions as a varifocal mirror for axial scanning. First, the system's focal shift capability was demonstrated with USAF resolution targets and carbon fiber phantoms, followed by in-vivo visualizations of blood vessels in chicken embryo chorioallantoic membrane (CAM). This work represents an initial step toward developing a non-contact, label-free, and aberration-free PARS imaging system with axial scanning capability.
© 2022 Optica Publishing Group under the terms of the Optica Open Access Publishing Agreement
1. Introduction
Photoacoustic microscopy (PAM) is a hybrid imaging technique that relies on optical absorption contrast and acoustic detection [1]. The excitation light energy absorbed by biomolecules is converted into heat, leading to thermoelastic expansion and the generation of acoustic waves [2]. Since acoustic waves scatter less than light in biological tissues, photoacoustic microscopy represents an attractive modality for high-resolution imaging at greater depths than purely optical approaches [3]. However, a major drawback of conventional photoacoustic microscopy is the requirement for an ultrasound transducer to be in direct physical contact with the sample through a coupling medium [4]. Such contact increases the risk of infection and contamination in open wound diagnostics and can result in microscopic injuries and patient discomfort in ophthalmic applications [5,6]. These complications make transducer-based photoacoustic techniques undesirable in many clinical and pre-clinical applications.
Photoacoustic remote sensing (PARS) microscopy is an all-optical, non-contact, label-free, and non-interferometric imaging technique to detect photoacoustic signals [7]. In PARS microscopy, a conventional ultrasonic transducer is replaced by an interrogation beam, co-aligned and co-focused with the excitation beam onto a sample. Due to thermo-elastic expansion, the energy absorbed by chromophores is converted to pressure. The increase in pressure creates elasto-optic modulations in the local optical properties within absorbers, thus altering the intensity of the back-reflected interrogation beam. PARS offers chromophore-specific contrast by targeting the unique absorption spectra of biomolecules with different excitation wavelengths. For example, DNA/RNA-containing structures are targeted by UV excitation light [8,9], while hemoglobin is targeted by visible light around 500 nm [10]. PARS microscopy has been successfully demonstrated in ophthalmic [11], H&E-like histology [12,13], blood vessels [14], and oxygen saturation imaging [15].
Conventional PARS differs from optical coherence tomography (OCT) in that it is not an interferometric technique, but PARS has an architecture to provide coherence-gated depth-resolved imaging [16]. Also, unlike PAM, PARS is not sensitive to photoacoustic time-of-flight. Instead, PARS detects photoacoustic pressures induced by pulsed lasers at their origin, so it is not an inherent 3D imaging system. Conventional PARS microscopy architectures have been optimized primarily to acquire two-dimensional data. Volumetric imaging is achieved by mechanical scanning [17,18]. The disadvantages to this method are slow mechanical scanning rates, vibrations and artifacts introduced during live imaging, as well as the bulky and expensive nature of the system [19]. An optical approach to shifting the focus may enable PARS to image larger volumes at higher speeds and at high resolution for multiple applications, including brain imaging [20], tumor angiogenesis [21,22], and oxygen saturation studies [3].
Optical axial scanning can be achieved by introducing a stationary active element called a varifocal element. Prime examples of varifocal elements include liquid-filled lenses, liquid crystals, vertical microlens scanners, and deformable mirrors (DMs) [23]. Most of them achieve the focal shift by changing their optical and/or geometric properties. MEMS DMs are the most common wavefront correctors due to their ease of manufacture, low cost, and high optical performance [24]. DMs are typically used as a part of adaptive optics (AO). This add-on technology adjusts the wavefront of the optical beam to compensate for system or sample-induced aberrations, improving the resolution of their respective imaging systems [25].
In this work, we have integrated a DM into a PARS microscope for microvasculature imaging at varying depths. The DM utilized in experiments is a novel continuous MEMS DM [26]. It features a simplified control mechanism and a resonant electrostatic actuation scheme that employs a single harmonic drive signal to replicate up to 8 low and high-order Zernike modes. This actuation methodology eliminates the need for individually addressable electrodes, complex control mechanisms, and associated hardware. Furthermore, the DM's stroke is increased by dynamic amplification, which occurs when the DM is driven at resonance [27].
In our experiments, we used the characteristics of the DM to create an optical model and predict the focal shift using Zemax. Then, the focus shifting ability of the DM was characterized experimentally by utilizing a 532 nm scattering microscope embedded in a PARS microscope. Subsequently, the axial scanning capabilities of DM-based PARS microscopy were demonstrated by imaging carbon fibers. Finally, in-vivo PARS imaging of blood vessels in chicken chorioallantoic membrane (CAM) models were performed at different depths by optically shifting the focal plane.
2. Methods
2.1 Resonant deformable mirror structure and operation
The MEMS DM [26] used in the experiments is made of a 1.6 mm diameter crystal silicon plate with a thickness of 10 µm (Fig. 1(A)) covered with a 75 nm thick gold layer to create a reflective surface. Eight equally spaced beams support the mirror plate. Electrostatic actuation drives the DM via 49 electrodes patterned in four concentric tiers on the bottom wafer 20 µm below the mirror plate. Fig. 1(B) shows two DMs on a chip carrier. One of them is wire-bonded and used in the experiment. The electric field is created by applying a single resonant harmonic voltage to one or more electrodes while the mirror plate is grounded. The operating principle of the DM depends on the use of a pulsed laser beam. The pulse repetition rate (PRR) of the laser is synchronized with the DM oscillations.
Fig. 1. (A) Schematic of the DM structure, (B) Two DMs under 5x magnification microscope, (C) Defocus mode obtained using a Laser Doppler Vibrometer.
The resonant DM can produce the eight lowest order Zernike modes, namely defocus, primary coma, astigmatism, primary spherical, trefoil, tetrafoil, secondary coma, and secondary spherical at different excitation frequencies and a single excitation signal. For example, the mirror is actuated at 8 kHz for a defocus mode and 40.5 kHz for a primary spherical mode. This study used the defocus mode, the first axisymmetric mode of the DM shown in Fig. 1(C), as a varifocal mirror for axial scanning. The mirror was actuated by applying a harmonic voltage waveform with an amplitude of 150 V and a frequency of 8 kHz to all actuation electrodes. Its radius of curvature was controlled by varying the phase angle ϕ between the pulse signal of the incident laser beam and the drive signal of the DM.
2.2 PARS system architecture
The PARS system illustrated in Fig. 2 is a modified version of the traditional PARS imaging setup outlined in [7]. The 532 nm 1.5 ns output of the fiber laser (GLPM-10, IPG Photonics) served as an excitation source for the imaging system. The output beam of the laser was coupled to an optical fiber (PM-460-HP, Thorlabs). The fiber output was coupled to a collimator mounted on a rotating holder to control the polarization. The beam size was reduced to ∼750 µm using a 5X Galilean beam condenser (BC) to effectively utilize the defocus mode of the DM. A polarizing beam splitter (PBS) transmitted ∼90% of the excitation light to the DM. The DM was adjusted to be orthogonal to the incident beam. The reflected light from the DM was diverted towards a collimating lens (L) through the same PBS. The location of the collimating lens was determined using Zemax simulation, and a detailed description is provided in Section 2.3. The detection light was delivered by an 830 nm superluminescent diode (SLD830S-A20, Thorlabs). The excitation and detection beams were combined using a dichroic mirror (DMSP605R) and directed toward the objective lens using a large-beam galvanometer scanning mirror (GVS012/M, Thorlabs). The two beams were co-focused onto the sample using a 20X 0.4 NA objective (MY20X-824, Thorlabs). Depending on the phase angle, the beam diameter of the excitation beam measured at the objective site varied from approximately 1 mm to 2 mm. As a result, the depth of focus of the excitation beam varied from approximately 163 µm to 40 µm. The depth of focus of the detection beam remained fixed at approximately 254 µm. The detection light reflected from the specimen was collected by the same objective and guided onto an avalanche photodiode (APD430A/M, Thorlabs). An additional photodiode (DET10A2, Thorlabs), PBS, and a QWP were incorporated into the excitation path to collect the back-scattered excitation light from the sample by implementing a 532 nm scattering microscope. The 532 nm scattering microscope provided information on the location of the excitation plane, allowing better alignment of the system. In Fig. 2, blue dashed lines illustrate wavefront control provided by the deformable mirror, while pink dashed lines demonstrate axial scanning. A point acquisition was acquired for each pixel and recorded by a high-speed digitizer connected to the two photodiodes in the system. All signal processing and image formation steps were performed in C/C++ and MATLAB environments.
Fig. 2. Simplified PARS optical setup. C: collimator, BC: beam condenser, PBS: polarized beam splitter, QWP: quarter-wave plate, DM: deformable mirror, L: collimating lens, Cond: condenser lens, M: mirror, Dich.: dichroic mirror, SF: spectral filter, GV: galvanometer, OL: objective lens.
2.3 Zemax optical model
Zemax was used to simulate an optical setup. A 532 nm collimated input beam of 750 µm was used to represent an ideally collimated incoming beam illuminating the DM, as shown in Fig. 3. The wavelength was set to 532 nm as the DM was only incorporated on the excitation path in the PARS system. The DM was represented by a standard surface type with a clear semi-diameter of 800 µm. The radii of curvatures of the DM varied from 220 mm to 380 mm based on previous experimental results [27] with a step size of 10 mm. The incoming light first passed through a PBS. A positive lens (LA1708-AB, Thorlabs) was used as a collimating lens. The location of the collimating lens was determined by fixing the radius of curvature of the DM at 350 mm and optimizing it to obtain the collimated beam out. Finally, the last element in the setup was a paraxial lens with a focal length of 10 mm used to focus the light. The distance between the PBS and the DM was based on the physical setup of the PARS system. The location of the focal plane corresponded to the distance (thickness in Zemax) after the paraxial lens was calculated using the thickness solve – marginal ray height.
2.4 Chicken chorioallantoic membrane (CAM) model preparation
CAM models were prepared in-house as cost-effective and easy-to-handle samples for in-vivo imaging of blood vessels and microvasculature. For that purpose, fertilized eggs from the White Leghorn breed were purchased from Frey’s Hatchery in St. Jacobs, ON, Canada. The fertilized eggs were then placed into the automatic rotating hatcher for 72 hours at 37°C and relative humidity of 65-70%. After 72 hours, the eggs were placed on their sides for a resting period of 60 minutes to allow the embryo to rise to the top of the egg. Meanwhile, a CAM container was prepared by placing a plastic film on the top of a plastic cup and pushed down to form a trough. The plastic film was secured with a rubber band following the protocol outlined by Naik et al. [28]. The plastic film was then sprayed with 70% Isopropyl Alcohol (IPA) before the cracked eggs were dropped into their respective CAM holders. A Dremel handheld saw attached to a vertical stand was used to drill into the surface of the eggshells with a circular cutting disc. Before drilling each egg, the disc and eggshells were sprayed and disinfected with 70% IPA. The eggs were then held sideways above the Dremel disc while maintaining the orientation achieved during the resting process. A long but shallow cut was made at the surface of the eggshell from the 90° mark to the 270° mark, keeping the yolk at the top and avoiding any rotational movements. Once the shallow cut was completed, the egg was placed down on the plastic film within the CAM holder and gently cracked by pressing it against the sides of the CAM holder. After the extraction process was complete, the ex-ovo CAM was covered with a perforated plastic film for ventilation and maintenance of proper humidity. The ex-ovo CAMs were then placed inside a plastic container holding 2 inches of warm water. The plastic containers with the ex-ovo CAMs were then placed in the incubator at 37.8°C and with a relative humidity of >68% up to the imaging day. Fig. 4 represents the ex-ovo CAM development from day 0 through day 12. This study used 9 to 12-days-old CAMs for blood vessel imaging.
3. Results
3.1 Optical and experimental axial scanning capability
As a first step, the axial scanning capability of the DM was approximated in Zemax. The distance between the focusing lens and the focal plane location was calculated by varying the radius of curvature of the DM. The focal length of the focusing lens is equal to 10 mm. When an incoming beam is perfectly collimated, the distance between the focusing lens and the location of the focal plane is, therefore, equal to 10 mm. As the curvature of the DM changed from 220 mm to 380 mm, the distance between the focusing lens and the location of the focal plane increased from 9.84 mm to 10.04 mm, as shown in Table 1. As a result, the total focal shift calculated by the Zemax simulation was 200 µm.
Table 1. The focal plane distances from the paraxial lens for different radii of curvatures
The focal shifting ability of the DM was validated experimentally by imaging a 1951 USAF resolution target. The images were acquired using a 532 nm scattering microscope setup. Fig. 5(A) shows images of a 1951 USAF resolution target from group 3, elements 5 and 6. The smallest element (group 3 element 6) has a resolution of 14.30 lp/mm, corresponding to a line spacing of ∼70 µm. The images were obtained at three different depths by moving the XYZ stage from 0 µm to 120 µm and 240 µm. The DM surface was deformed at each depth by varying the driving phase angle from 0° to 45° and 90°. The total number of images acquired was 9. Higher resolution images highlighted in black were obtained when the sample plane and the focal plane coincided. The edge spread function (ESF) was acquired along the black line for each image. The line spread function (LSF), the derivative of ESF, was plotted, and the full-width at half maximum (FWHM) values were calculated to obtain image resolution, as illustrated in Fig. 5(B). Fig. 5(C) demonstrates that at a depth of 0 µm, the FWHM increased from 4.5 µm to 11.7 µm as the phase angle changed from 0° to 45° to 90°, and the image resolution decreased, consequently. At a depth of 120 µm, the most resolved image was in the middle and had an FWHM value of 5 µm. Lastly, at a depth of 240 µm, the FWHM decreased from 10.22 µm to 4.47 µm, as the phase angle increased from 0° to 90°, respectively. The experimental results suggest that the total focal shift provided by the DM is 240 µm. Since the depth of focus of the detection beam is 254 µm, the system’s focus shifting ability is limited by the performance of the DM.
Fig. 5. (A) USAF resolution target imaging with a 532 nm scattering microscope. (B) The edge spread function (ESF) and line spread function (LSF) from raw data. (C) Full-width half maximum (FWHM) values at different depths.
It may be noted that the Zemax simulation and the experimental results differ by 40 µm of focal shift. This might result from using a paraxial lens in Zemax and an actual objective lens in the experiments. Ideally, a black box of the objective lens would be used instead of the paraxial lens to improve the simulation results. In addition, Zemax’s simulation used an ideally collimated beam, which rarely exists in reality. Moreover, while the distances between the optical components in Zemax are based on the experimental setup, they are not 100% precise. Lastly, the radii of curvature used in the Zemax optical design were obtained at 200 V [27], while the focal shifting experiments on the USAF targets were performed at 300 V of the actuation voltage. All the factors mentioned above need to be addressed for the simulation results to better match the experimental results.
3.2 Phantom imaging with PARS
Next, the DM-based PARS system’s ability to optically shift the focal plane was demonstrated on carbon fibers. The imaging setup shown in Fig. 6(A) is composed of placing a layer of carbon fibers on a glass slide, covering it with a cover glass, and then adding another layer of carbon fibers on top. As a result, we get two layers of carbon fibers arranged perpendicular to each other and separated by a cover glass. The thickness of the cover glass was measured to be 151 µm. The image of the top layer of carbon fibers was acquired at a 0° phase angle, Fig. 6(B). A further increase in phase angle to 60° allowed an image of the carbon fibers at the bottom layer to be obtained as the focal plane moved deeper into the sample, Fig. 6(C). Meanwhile, the mechanical stage remained stationary within the system. Shadows cast by the top layer of carbon fibers can be seen in Fig. 6(C), as indicated by white arrows. In Fig. 6(D), the top (green) and bottom layers (purple) of carbon fibers were reconstructed to create a single image with depth information. As demonstrated by the experiment, we were able to optically shift the focal plane to image two layers of carbon fibers separated by a 151 µm thick cover glass using the DM for wavefront control within the PARS microscope.
Fig. 6. Phantom imaging of carbon fibers with PARS system. (A) Sample setup for imaging. (B) Top layer carbon fibers imaged at 0 µm depth. (C) Bottom layer carbon fibers imaged at approximately 150 µm depth. White arrows indicate the shadow cast by the top layer of carbon fibers. (D) Overlayed images of carbon fibers at different depths.
3.3 DM-based photoacoustic remote sensing microscopy in in-vivo studies
To demonstrate the in-vivo capabilities of the system, the blood vasculature of CAMs was imaged, as shown in Fig. 7. The CAM models were placed in an ice bath during the imaging session to reduce motion artifacts. Images were acquired with 16 kHz pulse repetition rates of the excitation laser. As with phantom imaging, in-vivo imaging experiments were conducted by scanning the x-y plane at a fixed depth, then changing the phase angle to move the focal plane to a new depth. Acquisition time for one xy plane was approximately one minute. As the z-location of the focal plane increased with the increasing phase angle, the PARS microscope imaged blood vessels at deeper layers. At 0 µm, there were almost no blood vessels observed, except for a slight hint that can be seen in the lower part of the image indicated by white arrows. As the focal plane moved approximately 13 µm deeper into the sample, a blood vessel was imaged, and some capillary beds were captured, but the microscope's resolution was not enough to resolve them. It can also be noticed that the blood vessel was not ideally located on the imaging plane and its left side went deeper into the sample. By moving another 13 µm deeper into the sample, the whole blood vessel can be seen. At approximately 90 µm, the intensity decreased as the focus moved beyond the blood vessel. However, the blood vessels can still be observed at that depth, as the biomolecules still absorb the excitation light even when not at an ideal focal plane.
Fig. 7. In-vivo imaging of CAM models at different depths using PARS microscopy with DM. FOV is 660 × 660 µm2
In this work, we have demonstrated the ability of the PARS system to optically shift the focal plane using the DM on carbon fibers and chick embryo CAM blood vasculature. The technology opens the door to developing non-contact, label-free, and aberration-free optical systems with axial scanning capabilities for rapid, high-resolution volumetric imaging of biological tissues. For instance, an ideal high-resolution DM-based PARS system with rapid depth scanning capability would be able to image newly formed vessels deep within tumors that tend to be very small and exhibit very low signals in angiogenesis studies.
Future work will focus on improving the speed of the DM-based PARS system to provide rapid depth scanning by automating the phase angle change. The resolution of the system will also be enhanced by increasing the beam diameter and utilizing the full aperture of the objective lens. This will allow the system to visualize small structures such as capillary beds and red blood cells. A new set of radii of curvatures of the DM at 300 V of actuation voltage will be obtained and Zemax simulation will be performed. While no significant effect on SNR was observed in this study, a qualitative and quantitative analysis of the reflected detection beam sensitivity and the SNR will be performed in the future to better understand the DM's performance and its impact on the system parameters. Moreover, the ability to optically shift the focal plane will be used for correcting chromatic shift caused by wavelength changes in multiwavelength applications such as oxygen saturation studies. Other modes of the deformable mirror can be used for aberration correction applications in PARS. For example, the primary spherical mode of the DM can be used to correct spherical aberrations in ophthalmology applications.
4. Conclusions
For many applications, such as brain imaging, tumor angiogenesis, or multiwavelength studies, shifting the focal plane optically, rather than mechanically, may be a more suitable method for imaging large volumes at high speeds and with high resolution. This work demonstrates the integration of a DM with PARS microscopy to enable optical depth scanning. The defocus mode of the novel DM was used as a varifocal mirror to shift the focal plane. The focal shifting ability of the DM was first simulated in Zemax and then demonstrated experimentally on USAF targets. The results suggest that 240 µm of optical focal shift can be achieved, experimentally. The depth scanning ability of the PARS system was then demonstrated on carbon fibers. Two layers of carbon fibers separated by a 151 µm cover glass were imaged by optically moving the focal plane from the top layer to the bottom layer. Finally, the DM-based PARS microscope's axial scanning capability was validated in in-vivo imaging of blood vessels in chicken CAM models.
Funding
New Frontiers in Research Fund – Exploration (NFRFE-2019-01012); Natural Sciences and Engineering Research Council of Canada (DGECR-2019-00143, RGPIN2019-06134); Canada Foundation for Innovation (JELF #38000); Mitacs (IT13594); Centre for Bioengineering and Biotechnology, University of Waterloo (CBB Seed fund); University of Waterloo; illumiSonics (SRA #083181).
Acknowledgements
The authors would like to thank Hager Gaouda, Jean Flanagan, and Nancy Gibson for their continuous support and help with CAM sample preparations. The authors acknowledge funding from the University of Waterloo, NSERC Discovery grant, MITACS accelerator program, Canada Foundation for Innovation (CFI-JELF), Centre for Bioengineering and Biotechnology seed funding, New Frontiers in Research Fund –exploration, and research partnership support from illumiSonics Inc.
Disclosures
Authors P. Haji Reza and K. Bell have financial interests in illumiSonics Inc. that partially supported this work.
Data availability
Data underlying the results presented in this paper are not publicly available at this time but may be obtained from the authors upon reasonable request.
References
1. J. Yao and L. V Wang, “Photoacoustic microscopy,” Laser Photonics Rev. 7(5), 758–778 (2013). [CrossRef]
2. P. Beard, “Biomedical photoacoustic imaging,” Interface Focus. 1(4), 602–631 (2011). [CrossRef]
3. H. F. Zhang, K. Maslov, M.-L. Li, G. Stoica, and L. V. Wang, “In vivo volumetric imaging of subcutaneous microvasculature by photoacoustic microscopy,” Opt. Express 14(20), 9317–9323 (2006). [CrossRef]
4. Z. Hosseinaee, M. Le, K. Bell, and P. Haji Reza, “Towards non-contact photoacoustic imaging [review],” Photoacoustics 20, 100207 (2020). [CrossRef]
5. H. F. Zhang, C. A. Puliafito, and S. Jiao, “Photoacoustic Ophthalmoscopy for In Vivo Retinal Imaging: Current Status and Prospects,” Ophthalmic Surg. Lasers Imaging Retina 42(4), S106 (2011). [CrossRef]
6. A. de la Zerda, Y. M. Paulus, R. Teed, S. Bodapati, Y. Dollberg, B. T. Khuri-Yakub, M. S. Blumenkranz, D. M. Moshfeghi, and S. S. Gambhir, “Photoacoustic ocular imaging,” Opt. Lett. 35(3), 270 (2010). [CrossRef]
7. P. Hajireza, W. Shi, K. Bell, R. J. Paproski, and R. J. Zemp, “Non-interferometric photoacoustic remote sensing microscopy,” Light: Sci. Appl. 6(6), e16278 (2017). [CrossRef]
8. S. Abbasi, M. Le, B. Sonier, K. Bell, D. Dinakaran, G. Bigras, J. R. Mackey, and P. Haji Reza, “Chromophore selective multi-wavelength photoacoustic remote sensing of unstained human tissues,” Biomed. Opt. Express 10(11), 5461–5469 (2019). [CrossRef]
9. S. Abbasi, M. Le, B. Sonier, D. Dinakaran, G. Bigras, K. Bell, J. R. Mackey, and P. Haji Reza, “All-optical Reflection-mode Microscopic Histology of Unstained Human Tissues,” Sci. Rep. 9(1), 13392 (2019). [CrossRef]
10. K. Bell, L. Mukhangaliyeva, L. Khalili, and P. Haji Reza, “Hyperspectral absorption microscopy using photoacoustic remote sensing,” Opt. Express 29(15), 24338–24348 (2021). [CrossRef]
11. Z. Hosseinaee, N. Abbasi, N. Pellegrino, L. Khalili, L. Mukhangaliyeva, and P. Haji Reza, “Functional and structural ophthalmic imaging using noncontact multimodal photoacoustic remote sensing microscopy and optical coherence tomography,” Sci. Rep. 11(1), 11466 (2021). [CrossRef]
12. B. R. Ecclestone, K. Bell, S. Abbasi, D. Dinakaran, M. Taher, J. R. Mackey, and P. Haji Reza, “Histopathology for Mohs micrographic surgery with photoacoustic remote sensing microscopy,” Biomed. Opt. Express 12(1), 654–665 (2021). [CrossRef]
13. B. R. Ecclestone, S. Abbasi, K. Bell, D. Dinakaran, G. Bigras, J. R. Mackey, and P. Haji Reza, “Towards virtual biopsies of gastrointestinal tissues using photoacoustic remote sensing microscopy,” Quant. Imaging. Med. Surg. 11(3), 1070–1077 (2020). [CrossRef]
14. K. L. Bell, P. Haji Reza, and R. J. Zemp, “Real-time functional photoacoustic remote sensing microscopy,” Opt. Lett. 44(14), 3466–3469 (2019). [CrossRef]
15. Z. Hosseinaee, N. Pellegrino, N. Abbasi, T. Amiri, J. A. Tummon Simmons, P. Fieguth, and P. Haji Reza, “In-vivo functional and structural retinal imaging using multiwavelength photoacoustic remote sensing microscopy,” (123AD).
16. K. L. Bell, P. Hajireza, and R. J. Zemp, “Coherence-gated photoacoustic remote sensing microscopy,” Opt. Express 26(18), 23689–23704 (2018). [CrossRef]
17. S. Abbasi, K. Bell, B. Ecclestone, and P. Haji Reza, “Live feedback and 3D photoacoustic remote sensing,” Quant. Imaging. Med. Surg. 11(3), 1033–1045 (2020). [CrossRef]
18. B. R. Ecclestone, Z. Hosseinaee, N. Abbasi, K. Bell, D. Dinakaran, J. R. Mackey, and P. Haji Reza, “Three-dimensional virtual histology in unprocessed resected tissues with photoacoustic remote sensing (PARS) microscopy and optical coherence tomography (OCT),” Sci. Rep. 11(1), 13723 (2021). [CrossRef]
19. G. Lan, G. Li, and T. F. Mauger, “Design of high-performance adaptive objective lens with large optical depth scanning range for ultrabroad near infrared microscopic imaging,” Biomed. Opt. Express 6(9), 3362–3377 (2015). [CrossRef]
20. Y. Gong, C. Huang, J. Z. Li, B. F. Grewe, Y. Zhang, S. Eismann, and M. J. Schnitzer, “High-speed recording of neural spikes in awake mice and flies with a fluorescent voltage sensor,” Science 350(6266), 1361–1366 (2015). [CrossRef]
21. P. Carmeliet and R. K. Jain, “Angiogenesis in cancer and other diseases,” Nature 407(6801), 249–257 (2000). [CrossRef]
22. L. Nie, S. Wang, X. Wang, P. Rong, Y. Ma, G. Liu, P. Huang, G. Lu, and X. Chen, “In Vivo Volumetric Photoacoustic Molecular Angiography and Therapeutic Monitoring with Targeted Plasmonic Nanostars,” Small 10(8), 1585–1593 (2014). [CrossRef]
23. S. J. Lukes, “Dynamic and Agile Focusing in Microscopy: A Review,” https://doi.org/10.1117/3.2248542.ch1 SL20, 1–36 (2016).
24. T. Bifano, “MEMS deformable mirrors,” Nat. Photonics 5(1), 21–23 (2011). [CrossRef]
25. M. J. Booth, “Adaptive optics in microscopy,” Philos. Trans. R. Soc., A 365(1861), 2829–2843 (2007). [CrossRef]
26. S. Kocer, L. Mukhangaliyeva, R. Saritas, A. Gulsaran, A. Elhady, K. Bell, A. Kamel, M. Basha, T. Das, M. Kayaharman, P. Hajireza, M. Yavuz, and E. Abdel-Rahman, “Resonant Varifocal Mems Mirror,” in 2022 IEEE 35th International Conference on Micro Electro Mechanical Systems Conference (MEMS) (2022), pp. 963–966.
27. S. Kocer, “Resonant MEMS Deformable Mirror,” https://uwspace.uwaterloo.ca/handle/10012/17939.
28. M. Naik, P. Brahma, and M. Dixit, “A Cost-Effective and Efficient Chick Ex-Ovo CAM Assay Protocol to Assess Angiogenesis,” Methods Protoc. 1(2), 19 (2018). [CrossRef]
