MinHee K. Ko

Endothelial mesenchymal transformation mediated by IL-1β-induced FGF-2 in corneal endothelial cells.

This review describes the molecular mechanism of endothelial mesenchymal transformation (EMT) mediated by fibroblast growth factor-2 (FGF-2) in corneal endothelial cells (CECs). Corneal fibrosis is not frequently observed in corneal endothelium/Descemets membrane complex; but when this pathologic tissue is produced, it causes a loss of vision by physically blocking light transmittance. Herein, we will address the cellular activities of FGF-2 and its signaling pathways during the EMT process. Furthermore, we will discuss the role of inflammation on FGF-2-mediated EMT. Interleukin-1β (IL-1β) greatly upregulates FGF-2 production in CECs, thus leading to FGF-2-mediated EMT; the whole spectrum of the injury-mediated inflammation (IL-1β pathway) and the subsequent EMT process (FGF-2 pathway) will be briefly discussed. Intervention in the two pathways will provide the means to block EMT before inflammation causes an irreversible change, such as the production of retrocorneal fibrous membrane observed in human eyes.

The role of TLR4 activation in photoreceptor mitochondrial oxidative stress.

PURPOSE Herein the authors investigated whether the activation of Toll-like receptors (TLRs) in the innate immune response causes retinal photoreceptor oxidative stress and mitochondrial DNA (mtDNA) damage. METHODS On day 5 after injection of complete Freunds adjuvant containing heat-killed Mycobacterium tuberculosis (CFA), retinas were submitted to polymerase chain reaction (PCR) array focused on the TLR signaling, or apoptosis, pathway. CFA-mediated TLR4 activation, oxidative stress, and mtDNA damage were determined in B10.RIII and knockout (KO) mice (recombination activation gene [Rag] 1(KO), TLR4(KO), myeloid differentiation primary response gene 88 [MyD88](KO), tumor necrosis factor [TNF]-α(KO), or caspase 7(KO) mice) using quantitative real-time PCR, enzyme-linked immunosorbent assay, Western blot analysis, and immunohistochemistry. The mycobacterial DNA load on the retina, brain, liver, and spleen was determined by real-time PCR after intracardiac perfusion. RESULTS PCR array demonstrated the upregulation of TLRs and their signaling molecules in retinas of CFA-injected mice compared with those of control animals without inflammatory cell infiltration in the retina and uvea. Mycobacterial DNA was detected in the retinas of CFA-injected mice. Retinas of CFA-injected animals showed oxidative stress and mtDNA damage, primarily in the photoreceptor inner segments. Upregulated TLR4 was localized with CD11b(+)MHCII(+) cells but not with GFAP(+) astrocytes. This oxidative stress/damage was similar in CFA-injected Rag1(KO) mice compared with wild-type controls. Such damage was absent in the retinas of CFA-injected TLR4(KO), MyD88(KO), and TNF-α(KO) mice. CFA-mediated inducible nitric oxide synthase expression in the retina was significantly decreased in TNF-α(KO) mice. CONCLUSIONS Retinal photoreceptors are susceptible to mitochondrial oxidative stress/mtDNA damage in robust TLR4-mediated innate immune response.

Tissue-based multiphoton analysis of actomyosin and structural responses in human trabecular meshwork

The contractile trabecular meshwork (TM) modulates aqueous humor outflow resistance and intraocular pressure. The primary goal was to visualize and quantify human TM contractile state by analyzing actin polymerization (F-actin) by 2-photon excitation fluorescence imaging (TPEF) in situ. A secondary goal was to ascertain if structural extracellular matrix (ECM) configuration changed with contractility. Viable ex vivo human TM was incubated with latrunculin-A (Lat-A) or vehicle prior to Alexa-568-phalloidin labeling and TPEF. Quantitative image analysis was applied to 2-dimensional (2D) optical sections and 3D image reconstructions. After Lat-A exposure, (a) the F-actin network reorganized as aggregates; (b) F-actin-associated fluorescence intensity was reduced by 48.6% (mean; p = 0.007; n = 8); (c) F-actin 3D distribution was reduced by 68.9% (p = 0.040); (d) ECM pore cross-sectional area and volume were larger by 36% (p = 0.032) and 65% (p = 0.059) respectively and pores appeared more interconnected; (e) expression of type I collagen and elastin, key TM structural ECM proteins, were unaltered (p = 0.54); and (f) tissue viability was unchanged (p = 0.39) relative to vehicle controls. Thus Lat-A-induced reduction of actomyosin contractility was associated with TM porous expansion without evidence of reduced structural ECM protein expression or cellular viability. These important subcellular-level dynamics could be visualized and quantified within human tissue by TPEF.

Total Outflow Facility in Live C57BL/6 Mice of Different Age

Purpose: To characterize total outflow facility across the live adult mouse lifespan as a reference for mouse glaucoma studies and the common C57BL/6 background strain. Methods: Microperfusion was performed by single-needle cannulation and feedback-controlled coupling of pressure and flow to maintain a constant pressure in the anterior chambers of live C57BL/6NCrl mice aged 3-4 months (n = 17), 6-9 months (n = 10), and 23-27 months (n = 12). This mouse age range represented an equivalent human age range of young adult to elderly. We characterized the following across age groups in vivo: (1) outflow facility based on constant pressure perfusion in a pressure range of 15-35 mm Hg, (2) perfusion flow rates, and (3) anterior segment tissue histology after perfusion. Thirty-nine live mice underwent perfusion. Results: Pressure-flow rate functions were consistently linear for all age groups (all R2 > 0.96). Total outflow facility in mice aged 3-4, 6-9, and 23-27 months was 0.0066, 0.0064, and 0.0077 μL/min/mm Hg, respectively. Facility was not significantly different between age groups (all p > 0.4). The groups had closely overlapping frequency distribution profiles with right-sided tails. Post hoc estimates indicated that group facility differences of at least 50% would have been detectable, with this limit set mainly by inherent variability in the strain. A trend toward higher perfusion flow rates was seen in older mice aged 23-27 months, but this was not significantly different from that of mice aged 3-4 months or 6-9 months (p > 0.2). No histological disruption or difference in iridocorneal angle or drainage tissue structure was seen following perfusion in the different age groups. Conclusion: We did not find a significant difference in total outflow facility between different age groups across the live C57BL/6 mouse adult lifespan, agreeing with some human studies. The possibility that more subtle differences might exist ought to be judged with respect to the heterogeneity in facility at different ages. Our findings provide reference data for live perfusion studies pertaining to glaucoma involving the C57BL/6 strain.

Smooth muscle features of mouse extraocular muscle

We have been intrigued to find smooth muscle markers within mouse extraocular muscle (EOM) while studying contractile features of the aqueous humour drainage tissues by immunohistochemistry. Smooth muscle is known to be present in connective tissue fascial pulleys ensheathing EOM1 but not in the EOM per se. The mouse has many available engineered strains, shares many biological similarities with primates, and is widely used to model human biology, including that of EOM.2 Herein, we show that classic smooth muscle proteins of alpha-smooth muscle actin (α-SMA), myosin heavy chain (MHC), caldesmon, and tropomyosin are intimately associated with EOM fibre bundles, a finding that may be relevant to better understanding EOM control. We studied C57BL/6 mice (2-3 months, Charles River, Wilmington, MA) that were housed in a temperature-controlled room with 12h light and dark cycles and fed ad libitum. Animal care and use complied with the Institutional Animal Care and Use Committee as well as the Association for Research in Vision and Ophthalmology guidelines. Anterior (insertion) and posterior (retro-equator) portions of four rectus EOM of C57BL/6 mice (n=5) were carefully isolated, quickly embedded in OCT compound and snap-frozen in liquid nitrogen. Eight μm-thick cryosections were fixed with 4% paraformaldehyde, permeabilised and blocked (0.3% Triton X-100+5% BSA), incubated with primary antibodies (Abcam) to α-SMA, MHC non-muscle, caldesmon, and tropomyosin overnight at 4°C, then secondary antibodies and Alexa 568-phalloidin. Vascular smooth muscle labeling of the same tissues represented positive controls, while normal IgG isotype labeling represented negative controls. Sections were analysed by Leica SP5 or Zeiss LSM 710 confocal microscopy. The same tissue slides were further processed for hematoxylin and eosin staining to confirm structure. Figure 1 shows representative longitudinal and cross sectional immunohistochemistry images of the mouse anterior EOM at their globe insertions. F-actin labeling localized to contractile regions that were mostly consistent with striated muscle fibre bundles. Positive labeling for α-SMA, MHC, and caldesmon labeling was present in the periphery of phalloidin-positive muscle fibre bundles. Positive tropomyosin labeling was seen centrally and peripherally in muscle bundles, corresponding to regions of striated and presumed smooth muscle elements. Smooth muscle marker labeling partially overlapped with phalloidin labeling in fibre bundles. Figure 1 Smooth muscle profile in the anterior part of 4 different rectus extraocular muscles (EOMs). Co-localisation of α-SMA, MHC, caldesmon, or tropomyosin with phalloidin in the anterior portion of EOM. All 4 different rectus EOMs demonstrated a similar ... Figure 2 shows the distribution of smooth muscle markers in representative longitudinal and cross sections of rectus muscles posteriorly. All recti showed this pattern. As in the anterior EOM, α-SMA, MHC, and caldesmon labeling localized to the periphery of phalloidin-labeled bundles, while tropomyosin was present peripherally and centrally. Positive immunolabeling using the same prior antibody panel was seen in vascular smooth muscle from the same tissues (not shown). Figure 2 Smooth muscle profile of the posterior part of 4 different rectus EOMs. Co-localisation of α-SMA, MHC, caldesmon, or tropomyosin with phalloidin in the posterior portion of EOM was shown. All 4 different rectus EOMs demonstrated a similar in situ ... Orbital smooth muscle is present in superior and inferior palpebral muscles, inferior orbital fissure and EOM fascial pulleys1 as part of a periorbital smooth muscle network.5 To our best knowledge smooth muscle has not been described in EOM, which is considered striated muscle. Here we describe classic smooth muscle markers within mouse EOM, mostly in the periphery of striated muscle fiber bundles. Smooth muscle was present in all rectus muscles at their insertions and retro-equatorially, indicating this organization is widely present in mouse EOM. The EOM and their supporting structures represent a complex that helps orchestrate eye movement. Striated EOM are organized as global and orbital layers, each with different structural, vascular, neural, mechanical and metabolic features.3-4 Fascial sheaths and pulleys influence muscle actions on the globe1 under possible smooth muscle modulation. As with autonomically-innervated palpebral smooth muscle that works with striated muscle to regulate eyelid position,5 we postulate that smooth muscle associated with striated EOM plays a role in determining eye position in mice, and possibly in humans.