Table of Contents

Title Page

List of Contributors

Chapter 1: Sperm Selection Techniques and their Relevance to ART

1.1 Introduction
1.2 Need of Sperm Selection in ART
1.3 Methodology of Sperm Selection
1.4 Electrophoretic Sperm Separation
1.5 Zeta Test
1.6 Microelectrophoresis Sperm Selection
1.7 Raman Spectroscopy
1.8 Hyaluronic Acid Binding Assay
1.9 Future Perspective
References

Chapter 2: In Vitro Maturation of Human Oocytes: Current Practices and Future Promises

2.1 Introduction
2.2 Clinical Indications for IVM
2.3 Ovarian Stimulation Approaches for the Retrieval of Immature Oocytes
2.4 Maternal Conditions that may Influence IVM
2.5 Follicular Origins of Immature Oocytes for IVM
2.6 Clinical Safety of IVM
2.7 Concluding Remarks towards the Optimization of IVM
References

Chapter 3: Molecular Biology of Endometriosis

3.1 Introduction
3.2 Brief Background
3.3 Genetic Basis of Endometriosis
3.4 Molecular Mechanisms of Endometriosis
3.5 Molecular Etiopathological Basis of Endometriosis: Leads in Genomics Era
3.6 Molecular Etiopathological Basis of Endometriosis: Leads in the Post-Genomics Era
3.7 Future Targets
Acknowledgments
Conflicts of Interest
References

Chapter 4: Novel Immunological Aspects for the Treatment of Age-induced Ovarian and Testicular Infertility, Other Functional Diseases, and Early and Advanced Cancer Immunotherapy

4.1 Introduction
4.2 Ovarian Infertility
4.3 Novel In Vitro Proposals for Ovarian Infertility Treatment
4.4 Novel In Vivo Proposal for Ovarian and Testicular Infertility Treatment
4.5 Systemic Treatment of Other Functional Diseases by Tissue Rejuvenation
4.6 Advantages of Local and Systemic Use of Honey Bee Propolis and Cayenne Pepper
4.7 The Promise of Pyramid Healing Systems
4.8 Raw Shiitake Causes Early Neoplasia Regression and Malignancy Recurrence Prevention
4.9 Immune Modulation for the Treatment of an Advanced Cancer
4.10 Advanced Ovarian Cancer Regression Case Report
4.11 Discussion
4.12 Conclusions
Abbreviations
Competing Interests
Author Contribution
References

Chapter 5: Mitochondrial Manipulation for Infertility Treatment and Disease Prevention

5.1 Introduction
5.2 The Roles of Mitochondria in Fertilization, Embryonic Development, and Disease
5.3 The Genetics of Mitochondria and Mitochondrial Diseases
5.4 Ooplasmic Transfer to Treat Infertility
5.5 Pronuclear Transfer to Achieve Pregnancy
5.6 Germinal Vesicle Transfer to Restore the Viability of Oocytes
5.7 Mitochondrial Diseases and Prevention of their Inheritance
5.8 Mitochondrial Replacement by Transferring Pronuclei and MII Spindle
5.9 Discussion
Acknowledgments
References

Chapter 6: Novel Imaging Techniques to Assess Gametes and Preimplantation Embryos

6.1 Introduction
6.2 Light and Impact on Mammalian Gametes and Embryos
6.3 Novel Imaging Approaches for Gametes and Embryos
6.4 Conclusion
References

Chapter 7: Clinical Application of Methods to Select In Vitro Fertilized Embryos

7.1 Introduction
7.2 Morphological Assessment
7.3 Genomic and Transcriptomic Analysis
7.4 Analysis of Conditioned Culture Medium
7.5 Summary
References

Chapter 8: New Horizons/Developments in Time-Lapse Morphokinetic Analysis of Mammalian Embryos

8.1 Introduction
8.2 Utilization of Time-Lapse Morphokinetics in Mammalian Embryos: A Historical Perspective
8.3 What is TLM?
8.4 What are the Benefits of TLM?
8.5 Application of TLM in Human ART Practice
8.6 The Possible Utilization of TLM Analysis in Aneuploidy Detection
8.7 Expected Contributions of TLM Technology in the Future of Mammalian Embryology
References

Chapter 9: The Non-Human Primate Model for Early Human Development

9.1 Introduction
9.2 Why Primate Models Are Critical to Understanding Human Development and Subfertility
9.3 NHP Model of Assisted Reproductive Technology (ART)
9.4 NHP Model of Early Embryo Development
9.5 Research Perspective on NHP Embryo Development
9.6 Summary
References

Chapter 10: Cytoskeletal Functions, Defects, and Dysfunctions Affecting Human Fertilization and Embryo Development

10.1 Introduction
10.2 Components of the Cytoskeleton and their Important Functions in Reproductive Biology
10.3 The Role of the Cytoskeleton in Oocyte Maturation
10.4 Maturation Failures and Oocyte Aging
10.5 Fertilization and First Mitosis/Cell Division
10.6 Cellular Differentiation/Polarization During Pre-Implantation Embryo Development/Compaction Stage
10.7 Perspectives and Future Directions
References

Index

End User License Agreement

List of Illustrations

Chapter 1: Sperm Selection Techniques and their Relevance to ART

Figure 1.1 Apoptotic sperm are labeled by annexin V magnetic beads. A magnetic field separates the apoptotic sperm.
Figure 1.2 Separation of motile sperm using microfluidics.
Figure 1.3 Schematic diagram showing the apparatus for the electrophoretic sperm separation.
Figure 1.4 Diagram of sperm selection using the Zeta test. Negatively charged mature sperm is adhered to the positively charged tube surface, while immature sperm remain suspended in the media.
Figure 1.5 Schematic representation of microelectrophoresis unit.
Figure 1.6 A typical micro-Raman spectroscopy setup as utilized in the analysis of sperm.

Chapter 3: Molecular Biology of Endometriosis

Figure 3.1 Knowledge-based construction of the pathways-network of transcription factors putatively associated with the pathogenesis of endometriosis, obtained from functional analysis of co-expressed genes. CLOCK, ESR1, and MYC genes (shown inside blue dotted rectangle) appear to be involved in pathogenesis of endometriosis. Adapted from Ref. 154.
Figure 3.2 Pathways map derived from the enrichment analysis of differentially regulated genes between eutopic endometrium of subfertile patients with ovarian endometriosis in both proliferative and secretory phase of menstrual cycle highlighting important cellular processes getting involved in endometriosis. This Figure illustrates that molecular processes like angiogenesis (STAT signaling being the major mover), cell proliferation and apoptosis (MAPK signaling being the major mover) are affected in the eutopic endometrium of patients having ovarian endometriosis. Three major regulators – HIF1A, EGR1 and cFos (shown as asterisks) – are reportedly affected in endometroisis (142, 154, 160). Tables 3.3–3.5 highlight many of these genes in the process of development of endometriosis.
Figure 3.3 Cellular factors known to be associated with the pathways to neoplasm and tumorigenicity, some of which are reportedly dysregulated in eutopic and ectopic tissues in endometriosis as shown in red bullets. Interestingly, several studies on genomics expression, epigenetic changes and proteomics profile in endometriosis converge on the proposed involvement of these factors. For details, see Tables 3.3–3.7 and the text.
Figure 3.4 Immunopositivity of annexin A2 (a and b), HSP90 (c and d), PDGFRa (e and f), and Tubulin A (g and h) in eutopic endometria of subjects with diagnosed stage IV ovarian endometriosis during secretory phase of menstrual cycle showing their higher expression as compared to normal endometrium. Bars: 50 µm (c and d), 60 µm (h), and 75 µm (a, b, and e–g). Adapted from Ref. 196.
Figure 3.5 Venn diagram showing overlap of differentially expressed miRNAs in four different studies: Study A (244), Study B (278), Study C (279), and Study D (246). The four common miRNAs in these studies were miR-29c, miR-100, miR-200a, miR-200b. Some functional details of these miRNAs are given in Table 3.7. Adapted from Ref. 246.

Chapter 4: Novel Immunological Aspects for the Treatment of Age-induced Ovarian and Testicular Infertility, Other Functional Diseases, and Early and Advanced Cancer Immunotherapy

Figure 4.1 Immune type cells influence commitment of OSCs in adult human ovary (age 32 years, midfollicular phase). (a) Primitive CD14 MDC (green asterisk) associates with a small OSC (yellow asterisk and dotted circle) accompanying origination (green arrowhead) of a larger germ cell (red asterisk and dashed line) by asymmetric division of OSC (red arrowhead). (b) A serial section shows that asymmetric division is also accompanied by CD8 T cell (white asterisk) entering germ cell and exhibiting extensions (white arrowhead). (c) Divided primitive CD14 MDCs (green asterisks) accompany (green arrowheads) symmetric division (meiosis I cytokinesis) of germ cells (red asterisks) in the TA (ta) and germ cells moving (arrow) into the adjacent upper ovarian cortex (uoc). Inset shows a blood venule in the upper ovarian cortex with Thy-1 differentiation protein expression by vascular pericytes (arrow) and venule lumen (vl) containing a germ cell (red asterisk). (d) Germ cell transport in the upper ovarian cortex is associated with an attached activated (DR+) MDC (green asterisk and dotted lines) releasing DR+ cytoplasmic particles (green arrows) that accumulate at the surface of the germ cell nucleus (arrowhead). (e) Endothelial cells (en and open arrows) of a venule in the upper ovarian cortex exhibit MHC-I expression, which is not expressed by associated migrating (red arrow) germ cell (asterisk). See text for additional details. Adapted from Bukovsky et al. (1995b), with permission: © Blackwell Publishing, Oxford, UK.
Figure 4.2 Formation of fetal germ cells, granulosa cells, and follicular development in midpregnancy human fetal ovary. (a) CD14 MDC interacts (arrowheads) with an OSC (yellow asterisk and dotted circle) prior to the asymmetric division. (b) Numerous MHC-I depleted fetal germ cells (blue asterisks and dashed circles) originating by asymmetric divisions (arrowheads) from MHC-I+ OSCs (yellow asterisks and dotted circles). (c) Asymmetric division (blue arrowhead) is followed by a symmetric division of MHC I depleted germ cells (red asterisks and arrowhead). This is followed by the development of an ameboid shape moving germ cell (mgc – dashed line, no hematoxylin counterstain) entering adjacent ovarian cortex. Asymmetric division is accompanied by CD8 (d) and DR+ (e) T cell. (f) CD14 MDC (arrowhead) accompanies symmetric division of germ cell during meiosis I telophase. (g) Development of primitive granulosa cells (pgrc, note lower CK expression) from ovarian stem cells between mesenchymal cell cords (mcc). (h) DR+ MDC accompany (arrows) fetal growing follicle (gf) but not the resting follicle (rf). Inset shows association of Thy-1+ pericytes (arrowhead) with a growing but not resting follicles. Bar in a for a–f. Adapted from Bukovsky et al. (2005a) with permission: © Springer US.
Figure 4.3 Origin, meiosis I, and migration of human adult germ cells, and follicular renewal in a 28-year-old women (F28). (a) Dual color IHC of asymmetrically dividing OSC with CK+ (blue dashed line) OSC daughter and PS1+ (red dashed line) germ cell daughter. CK+ OSC daughter chromosomes (white arrowhead) move to the OSC end during mitotic OSC anaphase. Germ cell chromosomes (white arrows) duplicate by DNA replication (red and blue arrows) and exhibit sister chromatid crossover (orange arrows) during meiosis I prophase. White asterisk and dotted circle indicate PS1+ putative CD8/DR+ suicidal T cell within the germ cell (see Figure 4.1b, 4.2d, 4.2e and Bukovsky et al., 2001a). (b) In the TA (ta) the symmetrically dividing germ cell exhibits strong nuclear (asterisks) PS1 expression, which accompanies the meiosis I telophase. Arrow indicates a germ cell moving from the TA to the upper ovarian cortex (uoc). Inset shows a detail of chromosomal crossover (orange arrows) from panel A. Red and blue arrows indicate interacting sister chromatids. (c) Germ cell with a diminution of nuclear and increase of cytoplasmic PS1 staining. It begins to enter (arrowheads) the vein in the upper ovarian cortex. (d) Early stage of new primary follicle formation with ZP (blue color) expression of a small oocyte captured by the CK+ (brown color) granulosa cell nest. See details in the text. Adapted from Bukovsky et al. (2004): © Antonin Bukovsky.
Figure 4.4 Follicular renewal in adult human ovary and intravascular degeneration of germ cells unattended with granulosa cells. (a) Ovarian vein in the lower ovarian cortex lined by endothelial cells (en) and CK+ granulosa nest wall (gnw). In the vein lumen (vl) the granulosa nest wall extends a granulosa nest arm (gna) capturing the circulating oocyte (co). (b) Granulosa nest (gn) during formation of the new primary follicle with captured oocyte. Granulosa cells penetrate the ooplasm (red arrowheads) during the primary Balbiani body (asterisk) formation adjacent to the oocyte nucleus (on). CK+ granulosa nest particles (yellow arrowheads) are already dispersed within the oocyte, which still exhibits oocyte tail (ot) outside of the new primary follicle. (c) Growing preantral follicle (dashed line circle), with granulosa cells (grc) and oocyte (o) with ZP expression at the oocyte surface (arrow). (d) Degenerating oocyte in a medullary vein from the same ovary as in panel C exhibits a strong cytoplasmic ZP expression. (e) Heavily ZP+ degenerating oocyte from 28-year-old woman found in the extra ovarian (uterine ectocervix) vein of a patient with follicular renewal shown in panels a and b. Panels A and B adapted from Bukovsky et al. (2004): © Antonin Bukovsky; panels C–E adapted from Bukovsky et al. (2008b), with permission: © Elsevier/North-Holland Biomedical Press.
Figure 4.5 Cyclic formation of OSC, granulosa cells nests, and presence of new and resting primary follicles during midfollicular phase. (a) Tunica albuginea (ta) fibroblasts (fb) type OSC precursors with CK immunoexpression (brown). Two cells in mesenchymal-OSCs epithelial transition (fb/osc) are apparent. (b) Appearance of OSCs (osc) is associated with CK depletion (-fb). (c) Formation of CK+ granulosa cell nests is initiated by a layer of OSC (white arrows) above upper ovarian cortex (uoc). This is overgrown by a developing flap of TA (ta flap or taf in insert) resulting in a bi-layered osc cord (black arrow). Inset shows two layers of the OSC channel. (d) Detail of OSC flap with CK+ fibroblast type OSC precursors (fb/osc), and OSC development above the upper ovarian cortex (arched arrow). Arrowhead indicates the flap content of OSCs. (e) A parallel section to (D) showing numerous DR+ MDC (asterisks) in the TA flap. Note DR expression in early OSC (arrow). (f) Detail of OSC-cord from panel C shows CK+ epithelial cord. (g) OSC flap (red arrowhead) over a segment of TA (dashed line) covered by OSC layer (red arrow). The OSC cord-derived granulosa cell clusters (black arrows) fragment into granulosa cell nests (black arrowheads). Dashed line indicates a segment of TA covered by OSC epithelium. (h) Granulosa cell nests (black arrowheads) move by stromal rearrangements (arched arrow) to the lower ovarian cortex (loc) and form new primary follicles (white arrowhead) containing ZP+ oocytes. (i) Lower ovarian cortex (loc) with new primary (right panel segment) and resting primary follicles (left). Right inset shows the presence of primary Balbiani bodies. Left inset shows lack of Balbiani bodies. Bar in (a), for (a and b), bar in (f) for (d–f). Adapted from Bukovsky et al. (2004): © Antonin Bukovsky.
Figure 4.6 in vitro developing oocytes are supplied with meiotically nonfunctional organelles from fibroblasts or satellite cells. Time lapse photography shows that early developing oocytes (o, panel (a) are low in optically dense cytoplasmic organelles (white open arrow). They can be joined (arrowhead) by fibroblast-like cells (fb), providing additional organelles. Such fibroblast-type cells initially show optically dense organelles close to the nucleus (black solid arrow), but not in the arm extended toward the oocyte (white solid arrow). Within 10 min (panel (b), however, the optically dense organelles are apparent in the extended arm (solid black arrow) and within adjacent oocyte cytoplasm (black arrowhead) and distant oocyte regions (black open arrow). At 4h 25 min (panel c), however, the fibro-oocyte (fbo) hybrid is formed and regressing oocyte (ro) exhibits depletion of organelles (arrow) accumulated by the fibroblast (arrowhead). Alternatively, the developing oocytes (o, panel d) deficient in cytoplasmic organelles (white arrow) exploit the satellite cells (s), which are produced by the oocytes themselves. The oocyte is supplied by suicidal satellite cell by a tube like ring canal (black arrowhead; see inset). In panel e the oocyte exhibits enhanced content of the optically dense organelles (black arrow) and the ring canal draining the satellite disappears (white arrowhead – see inset). The satellite cell size is reduced (dashed line) and the perinuclear space is altered (compare with panel d). Oocytes enriched by satellites' organelles (panel f) exhibit good morphology [200 µm size, germinal vesicle (gv), and thick zona pellucida (zp)], but are unable to resume meiosis II due to the lack of meiotically functional organelles provided by secondary Balbiani body derived from granulosa cells in vivo. Bar in a for a–e. Panel c reprinted from Bukovsky and Caudle (2012): © Antonin Bukovsky. Other panels adapted from Bukovsky (2011b), with permission: © Wiley-Liss, Inc.
Figure 4.7 Time lapse video of oocyte reconstruction in secondary OSC culture (a) Early developing cell with a cytoplasmic tail (arrowhead). (b) Multiple cytoplasmic eruptions (arrowheads). (c) Development of the 50 µm oocyte-like cell (yellow arrowheads indicate cell surface, red arrowhead a polar body). Time in min':sec''. Reprinted from Bukovsky and Caudle (2012): © Antonin Bukovsky.
Figure 4.8 ZP and CK expression in ovarian follicles. (a) Oocytes in resting primary follicles lack ZP3 expression, but express ZP1, ZP2, and ZP4 (Bukovsky et al., 2008b). (b) ZP3 is expressed in the oocyte of a growing preantral follicle. (c) Double color IHC for CK (brown) and ZP (blue) expression in a growing preantral follicle. CK is expressed in granulosa cells (GrC) but no secondary Balbiani body is present in the oocyte. Oocyte surface expresses ZP (blue arrowhead). (d) Double color IHC for CK (blue) and ZP (brown) expression in a small antral follicle. CK is expressed in granulosa cells (GrC) and in the oocyte secondary Balbiani body (blue arrowhead). Oocyte surface expresses ZP (orange arrowhead).
Figure 4.9 ZP3 expression by OLCs in IVM treated OSC cultures is stolen by fibroblasts. (a) OLCs in untreated OSC cultures show week nuclear ZP3 expression, which is absent in accompanying satellite cells (SC) and fibroblasts (FB). Arrowheads indicate tube like ring canals between OLC and SC; arrows indicate bindings of FBs to the OLC. (b) After hCG treatment the OLC exhibits strong nuclear and surface (black arrowhead) ZP3 expression, which is also present in accompanying SC (white arrowhead). The ZP3 expression is stolen by FBs (red arrowheads), leaving the surface of OLC ZP3 depleted (open arrowhead). (c) The OSC culture from a 30-year-old POF female was IVM (FSH+hCG) pretreated and fertilized with the husband's sperm. The phase contrast (PhC) image from a live culture shows that the sperm are associated with fibroblasts instead with OLCs.
Figure 4.10 Propolis tincture preparation for the mouth and systemic use, and local effects for the hair, varicose veins, and teeth. (a) Measurement of 30 ml 40% alcoholic distillate. (b) Over layered propolis tincture for the local teeth treatment and dilution of the poured back propolis tincture for the systemic propolis treatment (c). (d) Developing frontal alopecia (arrow) before the local propolis treatment (back to front hair orientation) - note association of hair color depletion (dashed line circle) with developing alopecia. (e) Persisting unchanged frontal alopecia (arrow) (back to front hair orientation) – note restoration of the original hair color (compare with panel d) but persistence of color depletion in untreated coat sides (yellow arrowhead). (f) Hair condition in normal (side to side) hair orientation. (g) Recent hair appearance showing further improvement of hair condition, including color regeneration in propolis treated sides (red arrowhead). (h) Shrinking (blue arrows) and regressed (yellow arrows) varicose veins on the legs after the local propolis treatment. (i) Upper teeth row after professional dental cleaning and (j) heavy propolis deposits (arrows) thereafter on the densely brushed teeth. (k) Residual propolis attachments four days later. (l) Teeth status after 5 months of propolis treatment showing no propolis binding without any dental brushing, which indicates well-regenerated dental enamel. Arrowheads indicate a diminution of dental fissure (compare panel i). Numbers in blue rectangles indicate the image collection dates.
Figure 4.11 A Pyramid above the bed. Self-constructed pyramid consisting of 6.4 mm thick copper round, with a 53 cm long base and 49 cm long side cooper rounds. Reprinted from Bukovsky, 2016.

Chapter 5: Mitochondrial Manipulation for Infertility Treatment and Disease Prevention

Figure 5.1 The mitochondrial structure and functions. ATP is mainly produced through the oxidative phosphorylation (OXPHOS) pathway in mitochondria, where respiratory chain complexes I-IV act in coordination to create a proton gradient that drives ATP production by the complex V. The subunits of the respiratory chain complexes are encoded by nuclear and/or mitochondrial genes. Electrons flow from complex I and II to complex III through coenzyme Q (CoQ), from complex III to cytochrome c (Cyt. c), and from cytochrome c to complex IV, where four electrons and protons are used to reduce O₂ to H₂O. Although OXPHOS can leak electrons, which may generate free radicals, such as reactive oxygen species (ROS; O^·₂⁻), the free radical production is largely modulated by the rate of the electron flow.
Figure 5.2 The protocol for ooplasmic transfer.
Figure 5.3 Comparison of the three nuclear transfer protocols: germinal vesicle transfer, metaphase II spindle transfer and pronuclear transfer.

Chapter 6: Novel Imaging Techniques to Assess Gametes and Preimplantation Embryos

Figure 6.1 Representative images an (a) MII mouse oocyte with visualization of the meiotic spindle, (b) human sperm with birefringence and a vacuole, and (c) human oocyte zona pellucida showing three layers. (b) Adapted from Gianaroli et al. 2010, (c) adapted from Pelletier et al. 2004).
Figure 6.2 Representative images of rhesus oocytes with varying distribution of mitochondria (a, a', a') and (b). Rhesus 2PN zygote, (c) 1-cell, and (d) 2-cell embryo and mitochondrial distribution using Multiphoton image. Imaging required use of a fluoroprobe, but was compatible with continued embryo development (images adapted from Squirrell et al. 2003).
Figure 6.3 Representative images of a cleavage stage mouse embryo imaged with third (THG) harmonic generation (a). Image a' is the same embryo stained for lipids, which co-localized with the THG signal (a') (images adapted from Watanabe et al. 2010). Second harmonic generation imaging (SHG) can display structures such as the meiotic spindle (arrows; b, b', b') (images adapted from Thayil et al. 2011).
Figure 6.4 Representative images of a mouse GV-intact and MII oocyte imaged using Fourier transformed infrared. (a) GV intact oocyte and (a') MII oocyte; (b) chemical map of the integrated area under the amide bond of a GV oocyte and (B') MII oocyte; (c) CH₂ and CH₃ stretching region of a GV and (c') MII oocyte; (d) ester carbonyl band of a GV and (d') MII oocyte. IR imaging has been used to compare compositional differences, such as lipid distribution, between mouse oocytes of varying maturational status (images adapted from Wood et al. 2008).
Figure 6.5 Representative Raman microspectroscopy images of human sperm and a mouse oocyte. (a') brightfield image of human sperm and (a') chemical map constructed from Raman imaging of the same sperm using a cluster analysis from 400–3400 cm⁻¹, showing nucleus (green), neck (red) and midpiece (yellow). (b) Raman imaging of a mouse oocyte and chemical map constructed from cluster analysis from 500–1800 cm⁻¹ showing zona pellucida (green), subcortical (red) and central (pink) zone of cytoplasm. Raman imaging has been used to examine compositional content and changes in gamete DNA integrity and oxidative damage (images adapted from Bogliolo et al. 2013, Meister et al. 2010).
Figure 6.6 Representative CARS images of mouse GV-intact oocytes demonstrating lipid droplet distribution (images courtesy of J. Jasensky).
Figure 6.7 Representative images optical quadrature microscopy (OQM) of mouse embryos. OQM has been used to reliably count blastomeres compared to traditional staining/counting approaches (images adapted from Warger et al. 2007, Newmark et al. 2007).
Figure 6.8 Representative images of Optical Coherence Tomography (OCT) used to image mouse embryos before and after vitrification. Images A-C are unvitrified Metaphase I oocytes, 2PN and Metaphase II oocytes. Images D-F are images post-vitrification, where clumping of structures was observed and may be used to optimize the procedure. Identity of clumps is unknown (images courtesy of L. Zarnescu).
Figure 6.9 Representative image of phase subtraction microscopy of mouse embryos. Phase subtraction combines OQM and DIC microscopy and has been used to define blastomere boundaries and image embryos in the 3D plane, which may be used for cell counting. (images adapted from Warger et al. 2008).
Figure 6.10 Representative images of Qualitative Orientation Independent Microscopy (QOIM) of crane fly spermatocytes. QOIM uses DIC and polarization microscopy in tandem, and through the use of algorithms and computer-recombined images, produces a merged image (images adapted from Shribak et al. 2007, 2008).
Figure 6.11 Representative images of biodynamic imaging (BDI) of porcine (a) cumulus oocyte complexes and (b) blastocysts. Differences in cellular motions were detected between difference types of cells indicated a possible means of determining a non-invasive means of assessing subtle differences in cell quality (images adapted from An et al. 2015).

Chapter 7: Clinical Application of Methods to Select In Vitro Fertilized Embryos

Figure 7.1 Pre-implantation embryonic stages.

Chapter 9: The Non-Human Primate Model for Early Human Development

Figure 9.1 Induced DNA damage to sperm models, at least in part, human embryo developmental arrest. (a) control rhesus monkey ICSI embryo (no ROS treatment of sperm prior to fertilization), (b) ICSI embryo derived from ROS-treated motile sperm prior to fertilization of the oocyte, (c) phase contrast of normal four-cell ICSI rhesus embryo, (d) phase contrast of rhesus four-cell ICSI embryo displaying fragmentation of blastomeres, and (e) darkfield human embryo at the four-cell stage with fragments, a common arrest phenotype.
Figure 9.2 Rhesus embryo developmental landmarks for duration of cytokinesis in the first mitotic divisions after fertilization by intracytoplasmic sperm injection (ICSI).
Figure 9.3 Three rhesus blastocysts imaged using Eeva^TM and one arrested embryo (C1).

Chapter 10: Cytoskeletal Functions, Defects, and Dysfunctions Affecting Human Fertilization and Embryo Development

Figure 10.1 (a–d) From left to right: (a) MII meiotic spindle in unfertilized human oocyte is oriented perpendicularly to the oocyte surface. Insets: enlarged meiotic spindle and enlarged oocyte centrosome without centrioles (acentriolar centrosomes) of one meiotic pole. (b) Sperm incorporation. Inset shows sperm centrioles before fertilization. After fertilization the sperm aster forms from the proximal centriole of the centriole pair while the distal centriole disintegrates. (c) Zygote aster formation and two apposed pronuclei. (d) Mitotic apparatus of first embryonic cell division. Inset: enlarged mitotic apparatus and enlarged centrosome of one mitotic pole containing centrioles.
Figure 10.2 (a) Schematic diagram depicting stages of preimplantation human development. The zygote forms during Day 1. At the 8-cell stage (Day 3), embryos start to undergo compaction; the blastomeres become flattened and polarized. After the 8-cell stage, the inner cells are formed (Day 5: 32 cell stage) resulting in the inner cell mass (ICM); the outer cells form trophectoderm (TE). Days 6 to 9: Blastocyst formation and continued cell differentiation.
Figure 10.2

List of Tables

Chapter 3: Molecular Biology of Endometriosis

Table 3.1 Seven important cellular pathophysiological processes integral to the development of endometriosis
Table 3.2 Genes in pathophysiology of endometriosis based on candidate gene studies
Table 3.3 Candidate genes in endometriosis based on association studies
Table 3.4 Genes showing anomalous expression in endometriosis revealed from studies using customised large scale array-based transcriptomic studies
Table 3.5 Genome-wide transcriptomic studies revealing genes with affected expression in endometriosis
Table 3.6 Summary of published studies on proteomic profiles in eutopic endometrium in endometriosis
Table 3.7 Regulatory functions of a few differentially expressed miRNAs in endometriosis.¹
Table 3.8 Seven major regulatory processes associated with endometriosis based on molecular biology evidence
Table 3.9 Future therapeutic targets in endometriosis

Chapter 5: Mitochondrial Manipulation for Infertility Treatment and Disease Prevention

Table 5.1 The implementation and outcomes of ooplasmic transfer

Chapter 6: Novel Imaging Techniques to Assess Gametes and Preimplantation Embryos

Table 6.1 List of novel imaging approaches as well potential applications or novel information gained from gametes and embryos

Chapter 7: Clinical Application of Methods to Select In Vitro Fertilized Embryos

Table 7.1 Key time-lapse studies evaluating prediction of blastocyst development
Table 7.2 Key time-lapse studies evaluating implantation/pregnancy
Table 7.3 Studies evaluating aneuploidy
Table 7.4 List of metabolites in the literature, which have been correlated to implantation in human embryos. The regulation symbols are as follows: ↑ or ↓ denotes if the consumption of a metabolite is increased or decreased, respectively, in implanted embryos compared to non-implanted embryos, ÷ denotes that no differences were detected
Table 7.5 List of proteins in the literature, which have been identified in conditioned culture media from human embryos. The regulation symbols are as follows: ↑ or ↓ denotes if a protein is increased or decreased compared to control samples, + or ÷ denotes if the protein was detected in a positive or negative implantation group. * indicates that the observed regulation was not significant

Chapter 8: New Horizons/Developments in Time-Lapse Morphokinetic Analysis of Mammalian Embryos

Table 8.1 Different types of TLM systems that have been used in human embryo research in the literature
Table 8.2 Commercially available TLM systems that are currently available and used in human ART Laboratories
Table 8.3 Recent TLM studies and their correlation status with certain embryonic stages/events

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Title: Human reproduction : updates and new horizons / edited by Heide Schatten.

Description: Hoboken, New Jersey : John Wiley & Sons Inc., [2017] | Includes index.

Identifiers: LCCN 2016038683| ISBN 9781118849583 (cloth) | ISBN 9781118849576 (epub)

Classification: LCC QP251 .H845 2017 | DDC 612.6–dc23 LC record available at https://lccn.loc.gov/2016038683

List of Contributors

G. Anupa

Department of Physiology

All India Institute of Medical Sciences

New Delhi

India

Mustafa Bahceci

Bahceci Women's Health Group

Istanbul

Turkey

Muzaffer Ahmed Bhat

Department of Physiology

All India Institute of Medical Sciences

New Delhi

India

Antonin Bukovsky

The Laboratory of Reproductive Biology BIOCEV Institute of Biotechnology

Academy of Sciences of the Czech Republic

Prague

Czech Republic

Douglas T. Carrell

Andrology and IVF Laboratory

Department of Surgery (Urology)

Department of Obstetrics and Gynecology

and Department of Human Genetics

University of Utah

Salt Lake City

Utah

USA

Catherine M.H. Combelles

Biology Department

Middlebury College

Middlebury

Vermont

USA

Thomas F. Dyrlund

Department of Molecular Biology and Genetics

Aarhus University

Aarhus

Denmark

Necati Findikli

Bahceci Women's Health Group

Istanbul

Turkey

Debabrata Ghosh

Department of Physiology

All India Institute of Medical Sciences

New Delhi

India

Hans Jakob Ingerslev

The Fertility Clinic

Aarhus University Hospital

Aarhus

Denmark

Tetsuya Ishii

Office of Health and Safety

Hokkaido University

Hokkaido

Japan

Kirstine Kirkegaard

Department of Medical Biochemistry

Aarhus University Hospital

Aarhus

Denmark

Stuart Meyers

Department of Anatomy

Physiology

and Cell Biology

School of Veterinary Medicine

University of California

Davis

California

USA

Renee Riejo-Pera

Department of Cell Biology and Neurosciences

and Department of Chemistry and Biochemistry

Montana State University

Bozeman

Montana

USA

Heide Schatten

Department of Veterinary Pathobiology

University of Missouri

Columbia

Missouri

USA

Jayasree Sengupta

Department of Physiology

All India Institute of Medical Sciences

New Delhi

India

Munevver Serdaroğullari

Bahceci Women's Health Group

Istanbul

Turkey

Monis B. Shamsi

Andrology and IVF Laboratory

Department of Surgery (Urology)

University of Utah

Salt Lake City

Utah

USA

Luke Simon

Andrology and IVF Laboratory

Department of Surgery (Urology)

University of Utah

Salt Lake City

Utah

USA

Qing-Yuan Sun

State Key Laboratory of Reproductive Biology

Institute of Zoology

Chinese Academy of Sciences

Beijing

China

Jason E. Swain

Center for Reproductive Medicine

Department of Obstetrics & Gynecology

University of Michigan

Ann Arbor

Michigan

USA

Human Reproduction

Updates and New Horizons

List of Contributors