Open Access
Add to BibSonomyAbstract
The understanding of the reproductive events and the molecular mechanisms regulating fertility and infertility in humans relies heavily on the analysis of the corresponding phenotypes in mouse models. While molecular genetic approaches provide significant insight into the molecular regulation of these processes, the lack of live imaging methods that allow for detailed visualization of the mouse reproductive organs limits our investigations of dynamic events taking place during the ovulation, the fertilization and the pre-implantation stages of embryonic development. Here we introduce an in vivo three-dimensional imaging approach for visualizing the mouse oviduct and reproductive events with micro-scale spatial resolution using optical coherence tomography (OCT). This method relies on the natural tissue optical contrast and does not require the application of any contrast agents. For the first time, we present live high-resolution images of the internal structural features of the oviduct, as well as other reproductive organs and the oocytes surrounded by cumulus cells. These results provide the basis for a wide range of live dynamic studies focused on understanding fertility and infertility.
© 2015 Optical Society of America
1. Introduction and background
The mammalian oviduct is the conduit for oocytes ovulated by the ovary and the site of fertilization. Oviductal structure is thought to regulate sperm access to the ovulated oocytes for regulated fertilization. Subsequently, the fertilized oocyte transits through the oviduct, undergoing cleavage divisions for pre-implantation embryonic development and ultimately entrance into the uterus for implantation [1, 2]. In both humans and mice, it takes nearly four days for the oocytes to travel through the oviduct to the uterus [3, 4]. In the mouse, with a gestation of 20 days, this time frame covers a significant fraction of embryonic development. Defects associated with these processes can cause infertility [5–7], which motivated a wide range of studies investigating the causes and treatments of infertility from a molecular genetic perspective using the mouse as a model system [7–11]. However, the complete understanding of these processes from a dynamic perspective in their natural environment is missing, largely due to the lack of appropriate live high-resolution imaging methods in both humans and animal models [12–14].
A majority of studies focused on structural analysis of the mouse reproductive organs rely on histology [15, 16] or traditional optical bright-field microscopy and digital video-microscopy [15, 17, 18]. Histological techniques generally require the dissection, fixation and sectioning of reproductive organs, lacking the ability to capture dynamic processes. In contrast, traditional optical bright-field microscopy and digital video-microscopy are employed for direct visualization of the mouse oviduct and the live events of early embryogenesis, although these methods are hindered by a lack of depth-resolved information, a limited imaging depth and a low spatial resolution. Recently, electron microscopy and confocal florescence microscopy have been employed for the studies of the inner luminal surface of the mouse oviduct [19–21]. However, due to the limited imaging depth of these two techniques, the oviduct generally needs to be dissected from the animal and opened up to expose the luminal surface, potentially compromising in vivo investigations of tissue dynamics. Therefore, there is currently no imaging technique that allows for live depth-resolved high-resolution imaging of the reproductive events in the mouse oviduct.
To meet this demand, we developed and present an approach for in vivo volumetric high-speed imaging of the mouse oviduct and other reproductive organs with a resolution of ~5 μm using optical coherence tomography (OCT). OCT imaging was originally introduced in 1991 for noninvasive imaging of the retina [22]. Within the last two decades, OCT underwent rapid and dramatic developments with a major application in ophthalmology [23, 24], as well as some applications in oncology [25, 26], cardiology [27, 28], and developmental biology [29, 30]. The spatial resolution of modern OCT systems is ~1-15 μm, and the imaging depth in scattering tissues is around 1-3 mm [31, 32]. Previous studies have employed OCT for structural imaging of the in vitro dissected human fallopian tube and provided structural information comparable to histology [33, 34]. Very recently, Trottmann et al investigated the feasibility of using OCT for ex vivo imaging of the reproductive tract in the bovine model through a comparison with histology [35]. Computational approaches for analyzing the OCT images of the fallopian tube have also been developed for high-accuracy pathology diagnosis [36]. However, to the best of our knowledge, live OCT imaging of the mammalian oviduct have not been previously demonstrated.
In this paper, we present, for the first time, in vivo imaging of the internal structural details of the mouse oviduct using OCT. Reproductive organs were exposed for imaging in the anesthetized female through a small dorsal incision, similar to the traditional procedure of preimplantation embryo transfer during the production of genetically-modified mice. Live, high-resolution, three-dimensional visualizations of the developing follicles in the ovary, the oocytes surrounded by cumulus cells in the oviduct, as well as the unique structural features of the oviduct lumen are demonstrated. Our results suggest that OCT is a powerful imaging tool for in vivo mouse reproduction research, which opens the possibility for a wide range of live studies of reproductive events in mouse models.
2. Materials and methods
2.1 Mouse manipulation
CD-1 male and female mice were paired for overnight matings. The next morning, females were checked for the presence of a vaginal plug. The presence of the plug was counted as 0.5 days post conception (dpc). At the desired post-conception stage (0.5 dpc – 3.5 dpc), mice were anesthetized with a 1.25% tribromoethanol solution by intraperitoneal injection and placed on a heating pad to maintain body temperature at 37°C. Hair removal cream was applied to expose the dorsal lateral skin before the surgical procedure. The depilated skin was swabbed with 70% ethanol, a small incision (~1 cm) was made, and the reproductive organs (ovary, oviduct and a part of the uterine horn) were gently pulled out through the incision with blunt forceps and stabilized for imaging with a clamp. Tissue position was adjusted with forceps under a dissection microscope to orient the oviduct facing up (Fig. 1(A) and 1(B)). The animal was transferred to the imaging stage and the reproductive organs were imaged using OCT. The animals were euthanized after the imaging. All animal manipulation procedures were approved by the Animal Care and Use Committee of the Baylor College of Medicine.
Fig. 1 Experimental setup for in vivo OCT imaging of the mouse oviduct. (A) Mouse preparation with reproductive organs exposed and stabilized for in vivo OCT imaging. (B) Optical microscope image of mouse reproductive organs, specifically ovary, oviduct and uterus as oriented for OCT imaging. (C) A schematic of the imaging OCT setup with the spectral-domain configuration.
2.2 Spectral-domain OCT system
We utilized a home-built spectral-domain OCT system (Fig. 1(C)). The details of the OCT system have been described previously [37]. Briefly, the system employs a low-coherence Ti:Sapphire laser source (Micra-5, Coherence, Inc.) with a central wavelength of ~808 nm and a bandwidth of ~110 nm and an output power of ~400 mW. A fiber-based Michelson interferometer is utilized to produce the interference of the light backscattered from the sample and the light reflected from the reference arm. The interference fringes are spatially resolved with a home-built spectrometer, and fast Fourier transform is used to generate the OCT depth-resolved intensity A-lines. Two orthogonal galvanometer-mirrors are implemented in the sample arm for the transverse scanning of the laser beam across the sample, leading to three-dimensional volumetric imaging. A 5X scan lens is utilized to focus the beam on the sample. The OCT system has an axial resolution of ~5 μm in tissue and a lateral resolution of ~4 μm. Determined by the spatial sampling of the fringes, the available depth of imaging is measured around 5.5 mm in air, although due to the natural tissue scattering the imaging depth was about 1 mm. The in-depth pixel size is around 2 µm. The sensitivity of the system is measured to be ~97 dB for an optical path-length difference of around 50 µm with the exposure time of 18 µs. Over ~1 mm in depth, the sensitivity drop is ~4 dB. For in vivo mouse oviduct imaging, the A-scan acquisition rate is 50 kHz, and the 3D volume with 80 seconds data acquisition time can spatially cover a tissue area of ~6 mm × 6 mm in lateral directions. With 2000 × 2000 A-lines in one volume, it is densly sampled in the spatail dimensions with the transverse pixel size of 3 µm. For a smaller imaging region focusing on the specific parts of tissue, the lateral pixel scale is further improved. The 3D rendering and visualizations of the acquired OCT data is performed using Imaris software (Bitplane, Switzerland).
2.3 Histology
After the imaging, the animals were euthanized. The reproductive tissues were dissected, fixed in 4% methanol-free paraformaldehyde (PFA) for 2 hours at room temperature (RT), washed in phosphate-buffered saline (PBS) three times for 5 minutes, processed through sucrose gradient (10%, 20%, 30%) at 4°C ~3 hours each, then stored overnight in a 1:1 mix of 30% sucrose and optimal cutting temperature compound (O.C.T.), and frozen in O.C.T. media at −80°C. Cryosections of ~10 μm were fixed in 4% PFA for 5 minutes, washed 3 times in PBS (5 mintues each), permeabilized with PBT (1XPBS/0.1% Triton-X) 3 times (10 minutes each), washed 3 times in PBS (5 mintues each). For staining, sections were incubated with CF594 conjugated phalloidin (1:80 dilution) and diamidino-2-phenylindole (DAPI; 1:100 dilution) in PBS containing for one hour at RT. Slides were washed 3 times (5 minutes each) in PBS, mounted with flourmount-G and covered with micro coverslip glass. Sections were imaged with a fluorescent microscope using a 20x plan-apochromatic dry objective.
3. Results
Figure 2 shows a typical three-dimensional OCT reconstruction of the mouse reproductive organs, specifically the ovary, the oviduct, and the uterus. The oviduct is recognizable in Fig. 2(A) as the coiled structure located between the ovary and the uterus. Many structural features are clearly identifiable, such as the muscular striations of the uterus and the longitudinal folds of the oviduct running along the length of the oviduct. Figure 2(B) shows an in-depth cross-section of the reconstruction along the dashed line shown in Fig. 2(A). The inner lumen of the oviduct and the folded epithelium are visible, as well as the ovarian bursa which surrounds the ovary. While the whole depth of the oviduct was clearly resolved, OCT imaging provides only partial visualization of the uterus and the ovary due to high light attenuation in these tissues that restricts the imaging depth.
Fig. 2 OCT 3-D imaging of the mouse reproductive organs in vivo. (A) Three-dimensional OCT reconstruction of the mouse reproductive organs showing the external morphology of the ovary, the oviduct and the uterus. The dashed line represents the location for the cross-section shown in panel B. (B) Corresponding depth-resolved OCT cross-sectional image of the mouse reproductive organs showing the ovary, oviduct and uterus.
Mature oocytes develop postnatally from primordial follicles within the ovary under tight hormonal control. Follicular structure changes significantly as oocytes develop and mature prior to ovulation. Structural images of the ovary are shown in Fig. 3. The three-dimensional reconstruction of the exterior structure of the mouse ovary (ovarian bursa removed) with a portion of the oviduct are shown in Fig. 3(A), while the ovary inner structures are shown in Fig. 3(B) and Fig. 3(C). From the depth-resolved cross-sectional images (Fig. 3(B) and Fig. 3(C)), Graffian follicles with ova, less mature follicles and corpora lutea can be identified. Visualizing follicles in vivo at different developmental stages suggests that potentially the process of follicular development can be captured in longitudinal studies. Tearing the bursa (as well as the exposure of reproductive organs) is a part of a well-established procedure used for embryo transfer and production of transgenic mice [38]. This routine procedure does not compromise animal health and is followed by successful birth of pups. Therefore, it is not expected to affect the feasibility of longitudinal studies. While imaging of reproductive organs through the bursa is possible, tearing the bursa allows for higher imaging depth.
Fig. 3 OCT three-dimensional imaging of the mouse ovary in vivo. (A) Three-dimensional OCT reconstruction of the ovary with detailed inner structures visible. The dashed lines indicate the locations of the depth-resolved cross-sectional views of the ovarian tissue. (B) Depth-resolved OCT cross-sectional image of the mouse ovary highlighting a corpus luteum, a Graffian follicle with ova, as well as a less mature follicle. (C) An additional depth-resolved OCT cross-sectional image of the ovary containing the corpus luteum and a pre-ovulatory follicle. All scale bars correspond to 200 μm.
Upon ovulation, the oocytes surrounded by cumulus cells enter the oviduct through the infundibulum (the entrance to the oviduct adjacent to the ovary). After passing through the infundibulum, the ovulated oocytes enter the ampulla. The oviduct ampulla serves as the site of fertilization and transports the fertilized oocytes (zygotes) to the isthmus, the posterior portion of the oviduct. The detailed structural images from the ampulla of the mouse oviduct at 0.5 dpc acquired in vivo are shown in Fig. 4. The three-dimensional reconstruction (Fig. 4(A)) shows the longitudinal folds in the ampulla. Structural features such as height, orientation, and density of the folds can be visualized from the depth-resolved, cross-sectional images. (Fig. 4(B)) These features can potentially be used to define structural parameters and characterize oviduct tissue. A detailed view of the folds running through the lumen of the oviduct can be seen in Fig. 4(C). The cross-sectional views through the reconstruction capturing the oocytes inside the ampulla are shown in Figs. 4(D)-4(F). These images also visualize the zona pellucidae, an acellular membrane that surrounds each oocyte, and the surrounding cumulus cells. Despite sufficient spatial resolution and a high pixel density, the structural features within the oocyte and the cellular features of the oviduct cannot be resolved due to lack of optical contrast. However, the cumulus cells surrounding the oocytes in the ampulla are clearly visible. To the best of our knowledge this is the first in vivo three-dimensional visualization of the oocytes with cumulus cells within the oviduct, which provides the potential to study the dynamic activities occurring in the mouse reproductive tract.
Fig. 4 In vivo OCT imaging of the oviduct ampulla that contains the oocytes and the cumulus cells. (A) Three-dimensional OCT reconstruction of the structure of ampulla. Longitudinal mucosa folds are clearly visible in the image. The dashed lines indicate the locations of the depth-resolved cross-sections of ampulla. (B)-(F) OCT depth-resolved cross-sectional views of the ampulla showing the mucosa folds, the oocytes, zona pellucidae, and the cumulus cells in the oviduct of live mouse. All scale bars correspond to 100 μm.
After fertilization in the ampulla, cumulus cells detach from the zygote and begin their development and transit through the isthmus to arrive at the uterus for implantation around 4.0 dpc. The isthmus also serves as the site for cleavage divisions and embryo differentiation, resulting in the formation of blastocysts. Structural features of the oviduct as visualized with OCT are shown in Fig. 5. Figure 5(A) shows the highly coiled exterior structure of the oviduct isthmus, while Figs. 5(B)-5(F) show volumetric cross-sectional views through the same reconstruction exposing luminal space. A cross-section through three different folds of the oviduct with distinct folding patterns is shown in Fig. 5(B). In contrast to luminal longitudinal folds seen in the ampulla (Fig. 4), the folding pattern in the isthmus transitions to nodules of folded epithelium arranged in longitudinal rows in the anterior isthmus (solid arrow), followed by ring-like transverse folds in the posterior isthmus (dashed arrows). Figures 5(C)-5(F) provide additional cross-sectional views that highlight the nodules (solid arrow) and transverse (dashed arrows) folding patterns. The functions of these unique structures are currently unknown. These results indicate that OCT can be used to acquire high-resolution images of the oviduct isthmus in vivo and potentially be used to study the role of the oviduct folds in reproductive processes and analyze oocyte and cleavage-stage embryo development in their native environment.
Fig. 5 In vivo OCT imaging of the oviduct isthmus displays variable folding patterns. (A) Three-dimensional OCT reconstruction of the structure of the isthmus. The dashed lines indicate the locations of the depth-resolved cross-sections of oviduct. (B) Cross-sectional view of the inner lumen from the bottom surface. A solid red arrow indicates luminal, mucosal nodules arranged in longitudinal rows. Dashed arrows indicate ring-like transverse folding patterns. (C)-(F) OCT depth-resolved cross-sectional views of the oviduct showing the changes in mucosal structural patterns from longitudinal folds to nodules arranged in longitudinal rows, to transverse ring-like structures. All scale bars correspond to 200 μm.
Figure 6 shows comparison between in vivo OCT imaging of different structures of female reproductive tract and histological analysis of corresponding organs. There is a good correlation between structural features revealed through these two imaging approaches. Although the resolution of the fluorescence imaging is higher and provides subcellular structural details, our live OCT method is capable of acquiring comparable structural features in the imaged organs.
Fig. 6 Comparison of OCT images acquired in vivo to histological analysis of corresponding tissues. (A, C, E) Cross-sectional OCT images and (B, D, F) histology of the female reproductive tract. (A, B) Follicles in the ovary. (C, D) Images across the oviduct showing the folds in the lumen. (E, F) Cross-section along the oviduct showing the folds of the oviduct arranged in nodules. All scale bars correspond to 100 μm.
3. Discussions and conclusions
We present a novel imaging approach that allows for in vivo three-dimensional high-resolution visualization of the mouse reproductive organs and preimplantation events using OCT. The spatial resolution achievable with this approach is sufficient for visualizing detailed structural features of the ovary, oviduct, and oocytes with associated cumulus cells in a live mouse. The presented method has clear advantages over currently used methods for structural analysis of the mouse reproductive organs, specifically histology and confocal florescence microscopy. These advantages include the ability of in vivo imaging, a large field of view, and no requirement for exogenous contrast agents. Such features allow for dynamic investigations of the ovary, oviduct tissue and the oocytes with associated cumulus cells, with minimized alterations of their natural environments, which should facilitate advanced studies in female reproductive organs.
OCT imaging is considered to be safe and not damaging to tissue, as it has been widely used in ophthalmology and cardiology clinics [31]. Therefore, we do not expect any influence of the imaging light on the reproductive tissues. Compared with ultrasound imaging, one of the major advantages of using OCT for the imaging of mouse reproductive organs lies in the higher spatial resolution. OCT imaging clearly visualized the folds of the oviduct and the cumulus cells inside the oviduct lumen, which has not been possible using ultrasonic techniques, due to the resolution at tens or hundreds of microns. Currently, our OCT system provides a spatial resolution close to 5 μm in tissue. Employing broader bandwidth of the laser source could lead to single-micron OCT imaging of mouse oviduct [39], offering even higher spatial resolvability for cell-level imaging.
The OCT system used in this study employed near-infrared light that has a central wavelength around 808 nm. Since the attenuation of the light by the tissue is a major factor limiting the imaging depth, an OCT system with a longer central wavelength, such as around 1300 nm, could provide improved depth of view in the oviduct tissue [40, 41]. In addition, tissue optical clearing [42, 43] can potentially be applied in vivo to further enhance the OCT imaging of the deep structures.
The volumetric imaging speed is limited by the OCT A-line acquisition rate, which was set to 50 kHz in this study. According to our observations, such speed of data acquisition allows one to obtain densely sampled three-dimensional images of the mouse oviduct tissue in vivo without distortion or artifacts. To further improve the live imaging quality that might be affected by tissue motion (for example from the breathing of the mouse or muscle contractions), an OCT system with higher imaging speed could be implemented. For example, with a Fourier domain mode locking laser [44], an OCT A-line rate at the MHz level can be achieved [45], which will significantly reduce the time required to take the a three-dimensional volume of the oviduct. This could also benefit the imaging of dynamic events inside the female reproductive tract in the mouse model.
Although the procedure of exposing reproductive organs for manipulation through an incision in the body wall is well established, it is invasive. Implementing endoscopic OCT with the probe introduced to the reproductive organs through a small body wall incision might potentially provide a less invasive live imaging approach. Due to the limited imaging depth, the in vivo OCT imaging method presented here is intended as a research tool in the mouse model, and is not applicable for large animal models or human diagnosis. An endoscopic OCT probe with side-view imaging [46] could be a potential solution for in vivo, minimally invasive structural imaging of the human fallopian tube.
Currently, many mechanisms are still unknown regarding the differentiation and morphogenesis of the oviduct, such as the development of the folds [1]. One can improve the understanding of female infertility and develop advanced prevention and treatment protocols through investigations of the role of the oviduct in successful fertilization and embryo transit [5]. Since the mouse is an important mammalian reproductive model with numerous mutants available [1], the presented imaging method provides a powerful tool for a wide range of in vivo studies focused on reproductive and developmental biology in these models. Being able to successfully visualize structural features of the reproductive tract in live animals suggests the possibility for dynamic imaging and longitudinal studies using this approach.
To the best of our knowledge, the presented results are the first high-resolution in vivo three-dimensional visualizations of oocytes with cumulus cells in the mouse reproductive tract, which demonstrates the potential for embryo tracking and quantitative investigations of transport dynamics. Future work will focus on optimizing imaging methods for longitudinal studies as well as visualization and characterization of reproductive events during mammalian pre-implantation development.
Acknowledgments
Supported by grants from the National Institute of Health (R01HL120140, U54HG006348, T32GM008231) and the Optical Imaging and Vital Microscopy (OIVM) Core at Baylor College of Medicine.
References and links
1. C. A. Stewart and R. R. Behringer, “Mouse oviduct development,” Results Probl. Cell Differ. 55, 247–262 (2012). [PubMed]
2. J. E. Ellington, “The bovine oviduct and its role in reproduction: a review of the literature,” Cornell Vet. 81(3), 313–328 (1991). [PubMed]
3. H. B. Croxatto, “Physiology of gamete and embryo transport through the fallopian tube,” Reprod. Biomed. Online 4(2), 160–169 (2002). [CrossRef] [PubMed]
4. A. Bylander, M. Nutu, R. Wellander, M. Goksör, H. Billig, and D. G. J. Larsson, “Rapid effects of progesterone on ciliary beat frequency in the mouse fallopian tube,” Reprod. Biol. Endocrinol. 8(1), 48 (2010). [CrossRef] [PubMed]
5. R. H. F. Hunter, The Fallopian tubes: Their Role in Fertility and Infertility (Springer-Verlag, 1988).
6. M. M. Matzuk and D. J. Lamb, “The biology of infertility: research advances and clinical challenges,” Nat. Med. 14(11), 1197–1213 (2008). [CrossRef] [PubMed]
7. H. Wang and S. K. Dey, “Roadmap to embryo implantation: clues from mouse models,” Nat. Rev. Genet. 7(3), 185–199 (2006). [CrossRef] [PubMed]
8. R. Shao, J. Hu, and H. Billig, “Toward understanding chlamydia infection-induced infertility caused by dysfunctional oviducts,” J. Infect. Dis. 208(4), 707–709 (2013). [CrossRef] [PubMed]
9. M. Tuffrey, C. Woods, C. Inman, and M. Ward, “The effect of a single oral dose of azithromycin on chlamydial infertility and oviduct ultrastructure in mice,” J. Antimicrob. Chemother. 34(6), 989–999 (1994). [CrossRef] [PubMed]
10. R. E. Dixon, S. J. Hwang, G. W. Hennig, K. H. Ramsey, J. H. Schripsema, K. M. Sanders, and S. M. Ward, “Chlamydia infection causes loss of pacemaker cells and inhibits oocyte transport in the mouse oviduct,” Biol. Reprod. 80(4), 665–673 (2009). [CrossRef] [PubMed]
11. M.-W. Hu, Z.-B. Wang, H. Schatten, and Q.-Y. Sun, “New understandings on folliculogenesis/oogenesis regulation in mouse as revealed by conditional knockout,” J. Genet. Genomics 39(2), 61–68 (2012). [CrossRef] [PubMed]
12. M. A. Fritz, “The modern infertility evaluation,” Clin. Obstet. Gynecol. 55(3), 692–705 (2012). [CrossRef] [PubMed]
13. F. Petraglia, G. I. Serour, and C. Chapron, “The changing prevalence of infertility,” Int. J. Gynaecol. Obstet. 123(Suppl 2), S4–S8 (2013). [CrossRef] [PubMed]
14. S. Kölle, S. Dubielzig, S. Reese, A. Wehrend, P. König, and W. Kummer, “Ciliary transport, gamete interaction, and effects of the early embryo in the oviduct: ex vivo analyses using a new digital videomicroscopic system in the cow,” Biol. Reprod. 81(2), 267–274 (2009). [CrossRef] [PubMed]
15. G. Gonzalez and R. R. Behringer, “Dicer is required for female reproductive tract development and fertility in the mouse,” Mol. Reprod. Dev. 76(7), 678–688 (2009). [CrossRef] [PubMed]
16. S. Pal, W. Hui, E. M. Peterson, and L. M. de la Maza, “Factors influencing the induction of infertility in a mouse model of Chlamydia trachomatis ascending genital tract infection,” J. Med. Microbiol. 47(7), 599–605 (1998). [CrossRef] [PubMed]
17. S. S. Suarez, “Sperm transport and motility in the mouse oviduct: observations in situ,” Biol. Reprod. 36(1), 203–210 (1987). [CrossRef] [PubMed]
18. S. Kölle, S. Reese, and W. Kummer, “New aspects of gamete transport, fertilization, and embryonic development in the oviduct gained by means of live cell imaging,” Theriogenology 73(6), 786–795 (2010). [CrossRef] [PubMed]
19. D. Shi, K. Komatsu, M. Hirao, Y. Toyooka, H. Koyama, F. Tissir, A. M. Goffinet, T. Uemura, and T. Fujimori, “Celsr1 is required for the generation of polarity at multiple levels of the mouse oviduct,” Development 141(23), 4558–4568 (2014). [CrossRef] [PubMed]
20. M. Komatsu and H. Fujita, “Electron-microscopic studies on the development and aging of the oviduct epithelium of mice,” Anat. Embryol. (Berl.) 152(3), 243–259 (1978). [CrossRef] [PubMed]
21. M. C. Lee, T. C. Wu, Y. J. Wan, and I. Damjanov, “Pregnancy-related changes in the mouse oviduct and uterus revealed by differential binding of fluoresceinated lectins,” Histochemistry 79(3), 365–375 (1983). [CrossRef] [PubMed]
22. D. Huang, E. A. Swanson, C. P. Lin, J. S. Schuman, W. G. Stinson, W. Chang, M. R. Hee, T. Flotte, K. Gregory, C. A. Puliafito, and et, “Optical coherence tomography,” Science 254(5035), 1178–1181 (1991). [CrossRef] [PubMed]
23. W. Drexler, U. Morgner, R. K. Ghanta, F. X. Kärtner, J. S. Schuman, and J. G. Fujimoto, “Ultrahigh-resolution ophthalmic optical coherence tomography,” Nat. Med. 7(4), 502–507 (2001). [CrossRef] [PubMed]
24. L. An and R. K. Wang, “In vivo volumetric imaging of vascular perfusion within human retina and choroids with optical micro-angiography,” Opt. Express 16(15), 11438–11452 (2008). [CrossRef] [PubMed]
25. S. A. Boppart, W. Luo, D. L. Marks, and K. W. Singletary, “Optical coherence tomography: feasibility for basic research and image-guided surgery of breast cancer,” Breast Cancer Res. Treat. 84(2), 85–97 (2004). [CrossRef] [PubMed]
26. B. J. Vakoc, D. Fukumura, R. K. Jain, and B. E. Bouma, “Cancer imaging by optical coherence tomography: preclinical progress and clinical potential,” Nat. Rev. Cancer 12(5), 363–368 (2012). [CrossRef] [PubMed]
27. H. G. Bezerra, M. A. Costa, G. Guagliumi, A. M. Rollins, and D. I. Simon, “Intracoronary optical coherence tomography: a comprehensive review clinical and research applications,” JACC Cardiovasc. Interv. 2(11), 1035–1046 (2009). [CrossRef] [PubMed]
28. B. E. Bouma, G. J. Tearney, H. Yabushita, M. Shishkov, C. R. Kauffman, D. DeJoseph Gauthier, B. D. MacNeill, S. L. Houser, H. T. Aretz, E. F. Halpern, and I. K. Jang, “Evaluation of intracoronary stenting by intravascular optical coherence tomography,” Heart 89(3), 317–320 (2003). [CrossRef] [PubMed]
29. I. V. Larina, S. Ivers, S. Syed, M. E. Dickinson, and K. V. Larin, “Hemodynamic measurements from individual blood cells in early mammalian embryos with Doppler swept source OCT,” Opt. Lett. 34(7), 986–988 (2009). [CrossRef] [PubMed]
30. G. Karunamuni, S. Gu, Y. Q. Doughman, A. I. Noonan, A. M. Rollins, M. W. Jenkins, and M. Watanabe, “Using optical coherence tomography to rapidly phenotype and quantify congenital heart defects associated with prenatal alcohol exposure,” Dev. Dyn. 244, 607–618 (2014). [PubMed]
31. A. M. Zysk, F. T. Nguyen, A. L. Oldenburg, D. L. Marks, and S. A. Boppart, “Optical coherence tomography: a review of clinical development from bench to bedside,” J. Biomed. Opt. 12(5), 051403 (2007). [CrossRef] [PubMed]
32. J. G. Fujimoto, C. Pitris, S. A. Boppart, and M. E. Brezinski, “Optical coherence tomography: an emerging technology for biomedical imaging and optical biopsy,” Neoplasia 2(1-2), 9–25 (2000). [CrossRef] [PubMed]
33. J. M. Herrmann, M. E. Brezinski, B. E. Bouma, S. A. Boppart, C. Pitris, J. F. Southern, and J. G. Fujimoto, “Two- and three-dimensional high-resolution imaging of the human oviduct with optical coherence tomography,” Fertil. Steril. 70(1), 155–158 (1998). [CrossRef] [PubMed]
34. J. L. Fine, L. Kagemann, G. Wollstein, H. Ishikawa, and J. S. Schuman, “Direct Scanning of Pathology Specimens Using Spectral Domain Optical Coherence Tomography: A Pilot Study,” Ophthalmic Surg. Lasers Imaging 41(6), S58–S64 (2010). [CrossRef] [PubMed]
35. M. Trottmann, S. Kolle, R. Leeb, D. Doering, S. Reese, C. G. Stief, K. Dulohery, M. Leavy, J. Kuznetsova, C. Homann, and R. Sroka, “Ex vivo investigations on the potential of optical coherence tomography (OCT) as a diagnostic tool for reproductive medicine in a bovine model,” J. Biophotonics (2015).
36. M. Kirillin, O. Panteleeva, E. Yunusova, E. Donchenko, and N. Shakhova, “Criteria for pathology recognition in optical coherence tomography of fallopian tubes,” J. Biomed. Opt. 17(8), 081413 (2012). [CrossRef] [PubMed]
37. S. H. Syed, A. J. Coughlin, M. D. Garcia, S. Wang, J. L. West, K. V. Larin, and I. V. Larina, “Optical coherence tomography guided microinjections in live mouse embryos: high-resolution targeted manipulation for mouse embryonic research,” J. Biomed. Opt. 20(5), 051020 (2015). [CrossRef] [PubMed]
38. L. M. Ittner and J. Götz, “Pronuclear injection for the production of transgenic mice,” Nat. Protoc. 2(5), 1206–1215 (2007). [CrossRef] [PubMed]
39. L. Liu, J. A. Gardecki, S. K. Nadkarni, J. D. Toussaint, Y. Yagi, B. E. Bouma, and G. J. Tearney, “Imaging the Subcellular Structure of Human Coronary Atherosclerosis Using 1-μm Resolution Optical Coherence Tomography (μOCT),” Nat. Med. 17, 1010–1014 (2011). [CrossRef] [PubMed]
40. V. V. Tuchin, Tissue Optics: Light Scattering Methods and Instruments for Medical Diagnosis (SPIE press Bellingham, 2007), Vol. 642.
41. S. Ishida and N. Nishizawa, “Quantitative comparison of contrast and imaging depth of ultrahigh-resolution optical coherence tomography images in 800-1700 nm wavelength region,” Biomed. Opt. Express 3(2), 282–294 (2012). [CrossRef] [PubMed]
42. V. V. Tuchin, Optical Clearing of Tissues and Blood (SPIE Press Bellingham, 2006).
43. D. Zhu, K. V. Larin, Q. Luo, and V. V. Tuchin, “Recent progress in tissue optical clearing,” Laser Photon Rev 7(5), 732–757 (2013). [CrossRef] [PubMed]
44. R. Huber, M. Wojtkowski, and J. G. Fujimoto, “Fourier Domain Mode Locking (FDML): A new laser operating regime and applications for optical coherence tomography,” Opt. Express 14(8), 3225–3237 (2006). [CrossRef] [PubMed]
45. T. Klein, W. Wieser, L. Reznicek, A. Neubauer, A. Kampik, and R. Huber, “Multi-MHz retinal OCT,” Biomed. Opt. Express 4(10), 1890–1908 (2013). [CrossRef] [PubMed]
46. P. Herz, Y. Chen, A. Aguirre, J. Fujimoto, H. Mashimo, J. Schmitt, A. Koski, J. Goodnow, and C. Petersen, “Ultrahigh resolution optical biopsy with endoscopic optical coherence tomography,” Opt. Express 12(15), 3532–3542 (2004). [CrossRef] [PubMed]
