The levels of MDA and 8-iso-PGF2α, two classic oxidative products, in serum and spinal cord tissues were measured after SCI/R. The results showed that the levels of MDA (**Figure 4A**) and 8-iso-PGF2α (**Figure 4B**) in serum significantly increased after SCI/R in a time-dependent manner, but these increases were attenuated by NS309 treatment. The levels of MDA (**Figure 4C**) and 8-iso-PGF2α (**Figure 4D**) in spinal cord tissues from

FIGURE 4 | NS309 reduces the expression of oxidative products after SCI/R. The animals were treated with 2 mg/kg NS309 or vehicle at the beginning of SCI/R. The expression levels of MDA (A) and 8-iso-PGF2α (B) in serum were measured at different time points (0, 3, 6, 12, 24, and 48 h). The expression levels of MDA (C) and 8-iso-PGF2α (D) in spinal cord tissues were detected at 72 h after reperfusion. The data were represented as means ± SEM (n = 8). ∗p < 0.05 vs. Sham. #p < 0.05 vs. Vehicle.

NS309-treated animals was lower than that in vehicle group. In addition, we also detected the activities of SOD and CAT, two endogenous antioxidant enzymes. The results showed that the activities of SOD (**Figure 5A**) and CAT (**Figure 5B**) in serum from NS309-treated animals was higher than that in vehicle group from 12 to 48 h after SCI/R. As shown in **Figures 5C,D**, NS309 also markedly increased SOD and CAT activities in spinal cord tissues.

To investigate the effect of NS309 on inflammatory responses after SCI/R, the levels of inflammatory cytokines were determined in spinal cord tissues at different time points. The results showed that SCI/R significantly increased the levels of TNF-α (**Figure 6A**) and IL-1β (**Figure 6B**), two pro-inflammatory cytokines. However, application of NS309 significantly reduced the levels of these two cytokines in spinal cord tissues at 12, 24 and 48 h. Furthermore, we also measured the levels of IL-10 and TGF-β1, two anti-inflammatory cytokines. We found that NS309 markedly increased the levels of IL-10 at 24 and 48 h, but not at 12 h after SCI/R (**Figure 6C**). The TGF-β1 levels in NS309-treated group were higher than that in vehicle group at all time points measured (**Figure 6D**).

### NS309 Inhibits Mitochondrial Dysfunction After SCI/R

Mitochondrial function plays a key role in regulating oxidative stress under ischemia and reperfusion injury. Thus, we detected the activities of mitochondrial complexes in spinal cord tissues. The results showed that SCI/R significantly decreased the activities of all four mitochondrial complexes, and the activities of complex I (**Figure 7A**), complex III (**Figure 7C**) and complex IV (**Figure 7D**), but not complex II **Figure 7B**), were preserved by NS309 treatment. In addition, we investigated the effect of NS309 on mitochondrial biogenesis. NS309 increased mtDNA levels (**Figure 7E**) and elevated the levels of NRF-1 and PGC-1 mRNA after SCI/R (**Figure 7F**).

Mitochondrial dynamics are also involved in mitochondrial dysfunction under oxidative stress. As shown in **Figure 8A**, we

FIGURE 6 | NS309 regulates inflammatory cytokines after SCI/R. The animals were treated with 2 mg/kg NS309 or vehicle at the beginning of SCI/R. The levels of TNF-α (A), IL-1β (B), IL-10 (C), and TGF-β1 (D) in spinal cord tissues were determined at different time points (0, 12, 24, and 48 h). The data were represented as means ± SEM (n = 8). ∗p < 0.05 vs. Sham. #p < 0.05 vs. Vehicle.

detected the expression of mitochondrial dynamic proteins in spinal cord tissues after SCI/R at different time points. The results showed that SCI/R decreased the expression of Opa-1 and Mfn-1 at 12 and 24 h but increased the expression of Fis-1 from 3 to 24 h, with no effect on Drp-1 expression (**Figure 8B**). Furthermore, we detected the expression of these proteins after NS309 treatment (**Figure 8C**). We found that the decreased expression of Opa-1 and Mfn-1, as well as increased expression of Fis-1 after SCI/R, were all partially prevented by NS309 treatment (**Figure 8D**).

#### DISCUSSION

Due to the lack of data from randomized controlled clinical trials, contemporary concepts about SCI/R protection are mainly adapted from guidelines for brain ischemia, cardiovascular ischemia and spinal cord trauma (Nardone et al., 2016). This study provides evidence that NS309, a pharmacological activator for SK/KCa channel, protects against experimental SCI/R in rabbits. Our results showed that (a) SCI/R decreases SK/KCa1 protein expression in spinal cord tissues; (b) NS309 attenuates neuronal loss and neurological dysfunction after SCI/R; (c) NS309 reduces oxidative stress and regulates the levels of inflammatory cytokines after SCI/R; (d) NS309 preserves mitochondrial function and promotes mitochondrial biogenesis; and (e) NS309 differently regulates mitochondrial dynamic proteins.

The SK/KCa channels are extensively expressed throughout the central nervous system, but the three subtypes are differentially expressed. SK/KCa1 can be found in cortex, hippocampus, midbrain, cerebellum and spinal cord; SK/KCa2 is localized in cortex, amygdala, medial habenula and the inferior olivary complex; SK/KCa3 is expressed in olfactory bulbs, dentate gyrus thalamus and the locus coeruleus (Sailer et al., 2004). Thus, we detected the expression of SK/KCa1 protein to investigate the effect of SCI/R on SK/KCa channels. Significant decrease in SK/KCa1 expression was observed at 12 and 24 h after SCI/R, and this decrease was detected in both cytosolic and mitochondrial fractions. In neuronal cells, SK/KCa channels were originally thought to be predominantly presented in the plasma membrane where they regulates K+ efflux (Honrath et al., 2017). However, a previous study showed that SK/KCa channels were also located in the inner mitochondrial membrane of neuronal mitochondria (Dolga et al., 2013). In addition, Dolga et al. demonstrated that the SK/KCa2 subtype expressed in mitochondrial-enriched fractions and co-localized with the mitochondrial markers in neuronal dopaminergic cells (Dolga et al., 2014). Our results here demonstrated that SK/KCa1 was also localized in mitochondria in spinal cord, and was downregulated after SCI/R, indicating a potential role in mitochondrial function after SCI/R.

Activation of SK/KCa channels has been demonstrated to exert protective effects in many in vitro neuronal models, and is considered as an emerging therapeutic approach for the treatment of several neurological diseases (Anderson et al., 2006; Dolga et al., 2011, 2012, 2013, 2014; Dolga and Culmsee, 2012). NS309 is a selective and potent SK/KCa channel opener that activates SK/KCa channels via increasing their Ca2+ sensitivity (Strobaek et al., 2006). The EC50 of NS309 to SK/KCa channels is 30 nM, and it has been demonstrated to be safe and could activate SK/KCa channels in both in vitro and in vivo experimental models (Hipolito et al., 2015; Honrath et al., 2017). In this study, we found that NS309 treatment reduced spinal cord edema and spinal cord neuronal loss after SCI/R, which were accompanied by preserved neurological function. Our data extended the neuroprotective effects of SK/KCa channels activation into SCI/R. Importantly, we found that the protective effects of NS309 were still observed when it was administrated at 6 h after ischemia initiation. One of the serious problems of preclinical experiments is that many neuroprotective agents are administrated prior to injury initiation (Menon, 2009). It is of little clinical relevance due to the difficulty in obtaining informed consent. Our results showed that the therapeutic time window of NS309 was up to 6 h after ischemia initiation, which is useful for future clinical trials. This time window of administration makes it possible to treat SCI/R with NS309 associated neuroprotective agents. Considering the safety of NS309 treatment in previous experimental animal models, NS309 might be an ideal candidate for clinical drug research for SCI/R.

Free radicals-associated oxidative damage is one of the most important mechanisms underlying secondary injury after SCI/R, especially in the reperfusion phase (Chen X.P. et al., 2011). The overproduced ROS causes protein and lipid peroxidation, thereby leading to loss of membrane potential and corruption of membrane fluidity, which eventually results in the release of organelle content. In this study, increased levels of MDA and 8-iso-PGF2α, two oxidative products that correlate with the extent of ROS-mediated damage, was observed in both serum and spinal cord tissues after SCI/R, which were reduced by NS309. Neuronal fate under oxidative stress is dependent on the balance between ROS and the anti-oxidative defense mechanisms within the cell (Jia et al., 2012). SOD and CAT are two endogenous antioxidant enzymes, that can scavenge the superoxide radicals by changing the O2 − into less damaging species (Rodrigo et al., 2011). At 24 and 48 h after SCI/R, reduced activities of SOD and CAT was found to be preserved by NS309, indicating the anti-oxidative activity of NS309 in our model. Thus, the effects of NS309 on prevention of oxidative stress after SCI/R might be the direct result of the increased activity of antioxidant enzymes, which needs to be further determined in the future. Inflammation is a subsequent event of SCI/R, and inflammatory cell infiltration and release of inflammatory cytokines were observed in spinal cords (Zhu et al., 2013). The pro-inflammatory cytokines, such as TNFα and IL-1β, and anti-inflammatory cytokines, including IL-10

and TGF-β1, play opposite roles in inflammation regulation (Spitzbarth et al., 2012). Here, NS309 treatment was shown to upregulate anti-inflammatory cytokines but downregulate pro-inflammatory cytokines in the spinal cord tissues. Thus, inhibition of inflammatory responses via differentially regulating inflammatory cytokines might also contribute to NS309-induced protection against SCI/R.

Considering that the SK/KCa1 subtype was found in spinal cord mitochondria, and that NS309 alleviated oxidative stress after SCI/R, we speculated that activation of SK/KCa channels could regulate mitochondrial function in the spinal cord. Thus, we further measured mitochondrial complexes activity. It has been demonstrated that dysfunction of mitochondrial complexes occurred within 4–6 h after reperfusion and resulted in permanent mitochondrial damage (Sims and Muyderman, 2010). As expected, we found that the reduced activities of mitochondrial complexes after SCI/R, except for complex II, were all partially prevented by NS309. In addition, these effects were accompanied by the changes in mitochondrial biogenesis factors. Mitochondrial biogenesis is defined as the regulation of mitochondrial mass and turnover, a mechanism aimed to maintain diverse homeostatic demands under physiological and pathological conditions (Chen S.D. et al., 2011). It requires exquisite communication between the mitochondrial genome and the nuclear genome, and can be measured by the copy number and integrity of mtDNA, as well as the levels of many transcription factors, such as NRF-1, PGC-1, and TFAM (Kelly and Scarpulla, 2004). Previous studies have shown that these candidate mitochondrial biogenesis regulatory proteins correlate with mitochondrial respiratory capacity and could augment tolerance to ischemic injury via activating antioxidant system (McLeod et al., 2005). Thus, our results strongly indicate that NS309 might promote mitochondrial biogenesis, leading to inhibition of oxidative stress, and thereby induce protection against SCI/R.

Dynamic structural changes of the mitochondrial network, named as mitochondrial dynamics, are governed by the delicate balance between frequent fusion and fission events, and can adapt mitochondrial function to meet energy demand under stress (Otera et al., 2013). Impaired regulation of mitochondrial dynamic proteins contributes to ischemic injury via reducing energy production and promoting ROS generation (Calo et al., 2013). Multiple mitochondrial guanosine triphosphates hydrolases (GTPases) that regulate mitochondrial networking have been identified, among which DRP-1 and Fis-1 are related to mitochondrial fission, whereas mitochondrial fusion is mediated by Opa-1 and Mfn-1 (Karbowski et al., 2002; Hoppins et al., 2007). We found that SCI/R decreased the expression of Opa-1 and Mfn-1 but increased the levels of Fis-1 rather than Drp-1, indicating the induction of mitochondrial fragmentation in spinal cord after injury. In addition, the changes of these proteins after SCI/R were significantly prevented by NS309 treatment. Previous studies have shown that Opa-1 and Mfn-1 functionally interact to promote mitochondrial

elongation, while Fis-1 was required in the recruitment of Drp-1 to mitochondria during mitochondrial fission (Cipolat et al., 2004; De Palma et al., 2010). However, no significant changes in Drp-1 expression after SCI/R and/or NS309 treatment was observed in this study. Thus, a Drp-1-independent mechanism after SCI/R might exist, which need to be further determined.

In conclusion, our present study demonstrated the neuroprotective effects of NS309, a pharmacological activator for SK/KCa channel, against SCI/R in rabbits, which may involve an integrated process of the suppression of oxidative stress and the preservation of mitochondrial function.

#### REFERENCES

#### AUTHOR CONTRIBUTIONS

TC and Y-HW conceived and designed the experiments. JZ, L-KY, and W-LC performed the experiments. WL analyzed the data. TC and JZ wrote the paper.

## FUNDING

This work was supported by the National Natural Science Foundation of China (Nos. 81701932, 81601719, and 81871589).

in an inflammatory pain model. Pain 156, 849–858. doi: 10.1097/j.pain.<PHONE_NUMBER>000124

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Zhu, Yang, Chen, Lin, Wang and Chen. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Estradiol-Mediated Axogenesis of Hypothalamic Neurons Requires ERK1/2 and Ryanodine Receptors-Dependent Intracellular Ca2+ Rise in Male Rats

Lucas E. Cabrera Zapata1 , Mariana Bollo1 and María Julia Cambiasso1,2 \*

1 Instituto de Investigación Médica Mercedes y Martín Ferreyra, INIMEC-CONICET, Universidad Nacional de Córdoba, Córdoba, Argentina, 2Cátedra de Biología Celular, Facultad de Odontología, Universidad Nacional de Córdoba, Córdoba, Argentina

17β-estradiol (E2) induces axonal growth through extracellular signal-regulated kinase 1 and 2 (ERK1/2)-MAPK cascade in hypothalamic neurons of male rat embryos in vitro, but the mechanism that initiates these events is poorly understood. This study reports the intracellular Ca2+ increase that participates in the activation of ERK1/2 and axogenesis induced by E2. Hypothalamic neuron cultures were established from 16 day-old male rat embryos and fed with astroglia-conditioned media for 48 h. E2-induced ERK phosphorylation was completely abolished by a ryanodine receptor (RyR) inhibitor (ryanodine) and partially attenuated by an L-type voltage-gated Ca2+ channel (L-VGCC) blocker (nifedipine), an inositol-1,4,5-trisphosphate receptor (IP3R) inhibitor (2-APB), and a phospholipase C (PLC) inhibitor (U-73122). We also conducted Ca2+ imaging recording using primary cultured neurons. The results show that E2 rapidly induces an increase in cytosolic Ca2+, which often occurs in repetitive Ca2+ oscillations. This response was not observed in the absence of extracellular Ca2+ or with inhibitory ryanodine and was markedly reduced by nifedipine. E2-induced axonal growth was completely inhibited by ryanodine. In summary, the results suggest that Ca2+ mobilization from extracellular space as well as from the endoplasmic reticulum is necessary for E2-induced ERK1/2 activation and axogenesis. Understanding the mechanisms of brain estrogenic actions might contribute to develop novel estrogen-based therapies for neurodegenerative diseases.

Keywords: hypothalamic neurons, axogenesis, estradiol, ERK1/2, Ca2+ signaling, ryanodine receptors

## INTRODUCTION

For many years, estrogens have been recognized as one of the main orchestrators of the sexual differentiation of the brain, acting during critical periods of development to organize neural circuits in a way that determines the modulatory/activational effects of gonadal hormones in adulthood. Testosterone secreted by male rodent testes during development is aromatized in neurons to 17β-estradiol (E2), which displays multiple cellular processes that finally set the masculine phenotype (McCarthy, 2008; Wright et al., 2010). More recently, accumulating evidence

#### Edited by:

Gabriela Alejandra Salvador, Universidad Nacional del Sur, Argentina

#### Reviewed by:

Thomas Behnisch, Institutes of Brain Science, Fudan University, China Se-Young Choi, Seoul National University, South Korea

> \*Correspondence: María Julia Cambiasso

<EMAIL_ADDRESS>
Received: 15 January 2019 Accepted: 12 March 2019 Published: 02 April 2019

#### Citation:

Cabrera Zapata LE, Bollo M and Cambiasso MJ (2019) Estradiol-Mediated Axogenesis of Hypothalamic Neurons Requires ERK1/2 and Ryanodine Receptors-Dependent Intracellular Ca2+ Rise in Male Rats. Front. Cell. Neurosci. 13:122. doi: 10.3389/fncel.2019.00122

**88**

indicates that E2 is not only a reproductive hormone but also a brain-derived neuroprotective factor, coordinating multiple signaling mechanisms that protect the brain from neurodegenerative diseases, affective disorders and cognitive decline (Arevalo et al., 2015). These beneficial actions in the brain have positioned estrogens as promising therapeutic compounds against different brain pathologies such as Parkinson and Alzheimer diseases, schizophrenia, multiple sclerosis, stroke, neuroinflammation, among others (Dye et al., 2012; Villa et al., 2016; McGregor et al., 2017; Giatti et al., 2018).

Classical estrogenic actions are mediated via intracellular estrogen receptors (ERs) that function as ligand-activated transcription factors to regulate the expression of estrogenresponsive genes. Additionally, estrogens generate a wide diversity of rapid ''non-classical'' effects, which occur in a range from some seconds to a few minutes via membrane-initiated mechanisms (Arevalo et al., 2012), including the triggering of Ca2+ signals (Beyer and Raab, 1998; Picotto et al., 1999; Wong et al., 2012), and the activation of several signaling pathways, such as phospholipase C (PLC)/inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (Le Mellay et al., 1997; Chaban et al., 2004), nitric oxide synthase/nitric oxide (Kelly and Levin, 2001), adenylate cyclase/AMPc/protein kinase A (PKA; Beyer and Karolczak, 2000), phosphoinositide-3 kinase (PI3K; Garcia-Segura et al., 2010), PKC, and extracellular signal-regulated kinase 1 and 2 (ERK1/2) cascades (Wu et al., 2005).

 The induction of neurite outgrowth by E2 was first demonstrated by Toran-Allerand (1976, 1980) and Toran-Allerand et al. (1983) working with organotypic explant cultures of the preoptic area, hypothalamus, and cerebral cortex. This neuritogenic effect of the hormone was then observed in other brain regions, both directly related and unrelated with reproduction (Nishizuka and Arai, 1981; Reisert et al., 1987; Cambiasso et al., 1995; Murphy and Segal, 1996). Hypothalamic neurons in vitro undergo several intermediate stages of development from unpolarized to fully polarized cells (Díaz et al., 1992). Most of the neuritogenic effects of E2 were demonstrated in polarized neurons (stage III of development), which are characterized by the presence of axon (Díaz et al., 1992; Cambiasso et al., 1995). Previous studies from our laboratory have shown that E2 induces axonal growth through ERK1/2 activation in hypothalamic neurons of male embryos in vitro. Both axogenesis (Cambiasso and Carrer, 2001) and ERK1/2 activation (Gorosito and Cambiasso, 2008) mediated by the hormone are dependent on a membraneinitiated mechanism since E2-bovine serum albumin (BSA; a membrane-impermeable conjugate of E2) was as effective as free E2. Interestingly, decreasing intracellular Ca2+ by the Ca2+-chelator BAPTA-AM or blocking Ca2+-dependent PKC isoforms by Ro 32-0432 significantly decreased these E2 effects (Gorosito and Cambiasso, 2008). These findings strongly suggested an important role for Ca2+ in E2-induced ERK1/2 pathway activation and axonal growth; however, the results did not provide the mechanism of E2-induced Ca2+ signaling in hypothalamic neurons.

In this study, we found that E2 evoked activation of Ca2+ entry via L-type voltage-gated Ca2+ channels (L-VGCCs) and promoted Ca2+ release through ryanodine receptors (RyRs). This early Ca2+ response underlies E2-induced ERK1/2 activation and axogenesis in hypothalamic neurons. Altogether, these results bring new insights about the mechanism of brain estrogenic actions and might contribute to developing novel estrogen-based therapies for neurodegenerative diseases.

#### MATERIALS AND METHODS

#### Animals and Cell Cultures

Embryos were obtained from pregnant Wistar rats at embryonic day 16 (E16). The day of vaginal plug was set as E0. Experimental procedures for handling and sacrificing animals were approved by the Animal Care and Use Committee at our institution (CICUAL-IMMF, INIMEC-CONICET-UNC; Córdoba, Argentina) and followed the NIH guidelines for care and use of laboratory animals. The minimum number of animals required was used for these experiments and suffering was minimized. Primary neuronal and astroglial cultures were prepared as previously described in Cambiasso et al. (2000). Pregnant rats were sacrificed by cervical dislocation under CO2 anesthesia, and the fetuses were dissected from the uterus. The male fetuses used for cultures were identified by visualization of the spermatic artery on the developing testes. Ventromedial hypothalamic and mesencephalic regions were dissected out and stripped off the meninges for primary neuronal and glial cultures, respectively. At E16, the axogenic effect of E2 is contingent on the presence of astroglia (Cambiasso et al., 1995) or astroglia-conditioned media from a target region (Cambiasso et al., 2000; Cambiasso and Carrer, 2001; Brito et al., 2004). The basal medium (BM) was (1:1) DMEM:Ham's F12 Nutrient Mixture, supplemented with 0.043% l-alanyl-lglutamine (GlutaMAX I), 0.15% glucose, 100 U/ml penicillin and 100 µg/ml streptomycin. All cultures were raised under phenol red-free conditions to avoid ''estrogen-like effects'' (Berthois et al., 1986). For neuronal cultures, the dissociated cell suspension was seeded on different supports pre-coated with 1 mg/ml poly-D-lysine depending on the experiment: 60 mm × 15 mm dishes (Corning Life Science, Tewksbury, MA, USA) for protein assays, 25 mm coverslips (Assistent, Germany) for Ca2+ imaging, and 12 mm coverslips (Assistent, Germany) for morphological studies.

#### Western Blot

Hypothalamic neurons derived from male fetuses plated 1–2 h before were fed with astroglia-conditioned media for 2 days in vitro (DIV). After a 2 h washout period using BM, neuronal cultures were treated for 1 h with nifedipine (2 µM; Sigma-Aldrich, St. Louis, MO, USA), inhibitory ryanodine (50 µM; Santa Cruz Biotechnology, Santa Cruz, CA, USA), 2-APB (100 µM; Santa Cruz Biotechnology, Santa Cruz, CA, USA) or U-73122 (10 µM; Sigma-Aldrich, St. Louis, MO, USA), and then pulsed with 100 nM E2 (Sigma-Aldrich, St. Louis, MO, USA) for 15 min. Hormone concentration used was determined by dose dependence (1–100 nM) experiments previously performed by our group (Gorosito and Cambiasso, 2008; Gorosito et al., 2008). ERK phosphorylation was maximally increased after the application of 100 nM E2. This dose was then used for all further acute stimulation studies. We have used compounds at final concentrations that did not alter cell viability or morphology in control conditions.

After treatment, hypothalamic neurons were washed and harvested at 4◦C in RIPA buffer [150 mM NaCl, 0.1% NP40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate (SDS), 50 mM Tris, pH 7.5] with protease and phosphatase inhibitors (1 µg/ml aprotinin, 1 µg/ml leupeptin, 1 µg/ml pepstatin A, 5 µg/ml chymostatin, 5 µg/ml antipain, 100 µg/ml PMSF, 50 µM NaF, 10 µM Na4P2O7, and 1 mM NaVO4). Protein samples (20 µg/lane) were separated by 10% SDS-PAGE and transferred onto polyvinylidene fluoride membrane (Bio-Rad, Hercules, CA, USA). ERK1/2 phosphorylation was detected by using a rabbit monoclonal anti-phospho-p44/42 MAPK (Cell Signaling Technology, Danvers, MA, USA), which specifically detects both Thr202 and Tyr204 ERK phosphorylation forms of ERK [molecular masses (kDa) for ERK1 and ERK2 are 44 and 42, respectively]. Total ERK1/2 was detected by using a mouse anti-p44/42 MAPK (Cell Signaling Technology, Danvers, MA, USA). Secondary antibodies conjugated to horseradish peroxidase (Jackson, West Grove, PA, USA) were used for the detection by enhanced chemiluminescence on X-ray film. After incubation with the antibody against phospho-ERK1/2, blots were stripped and then re-probed with anti-total ERK1/2 to ensure equal protein loading. The resulting film samples were scanned and analyzed with an image analysis program (ImageJ; NIH, Bethesda, MD, USA). Data are presented as a ratio of phospho-ERK1/2/total-ERK1/2 of 3–4 different experiments (independent cultures) performed in duplicate.

### Ca2+ Imaging

After 2 DIV, hypothalamic neurons were treated (or not) with 50 µM ryanodine or 2 µM nifedipine for 1 h and incubated with 3 µM acetoxymethyl (AM) ester form of the organic Ca2+-dye Cal-520 (AAT Bioquest, Sunnyvale, CA, USA) for 30 min at 37◦C in a Ca2+-containing HEPES buffered salt solution (Ca2+- HBSS) composed of (mM): 135 NaCl, 5.4 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose; pH = 7.4 set with NaOH at RT. Following loading, neurons were washed twice with warm Ca2+- HBSS and were imaged in the same buffer. To analyze the participation of extracellular Ca2+, the cells were resuspended in an EGTA-containing buffer composed of (mM): 135 NaCl, 5 KCl, 0.5 CaCl2, 1.2 MgCl2, 5 HEPES, 14 NaHCO3, and 1 EGTA; pH = 7.4 set with NaOH at RT.

Imaging of cytosolic Ca2+ signals was performed using a 60× oil immersion objective of an Olympus IX81 inverted microscope [equipped with a Disk Spinning Unit (DSU), epifluorescence illumination (150 W Xenon Lamp), and a microprocessor], an ORCA AG (Hamamatsu) CCD camera and OSIS software. Frames were collected at a continuous rate of 2.5 per second during 5 min (790 frames). Cal-520 was excited at a wavelength of 492 nm, and emitted fluorescence was collected at 514 nm. E2 (100 nM) was added 30 s after starting the recording. 10 µM thapsigargin (tg) was added at 3 min of recording as a positive control of normal endoplasmic reticulum Ca2+ content. The fluorescence intensity of the Ca2+ indicator was analyzed using ImageJ (NIH, Bethesda, MD, USA) software and plotted as the change in fluorescence (∆F) of 2 × 2 pixels divided by mean resting fluorescence [(Fo; ∆F/Fo)] over time. We measured both the peak fluorescence value and the integrated area under the ∆F/Fo curve with OriginPro 8 SR0 software (OriginLab Corporation, Northampton, MA, USA). The integrated area roughly corresponds to the total amount of Ca2+ released over the recording period.

#### Immunocytochemical Staining

To analyze the effect of ryanodine in E2-stimulated axon growth without affecting the normal polarization of neurons, we performed the experiment after 1 DIV (stage III of development). After 2 h in absence of E2, the cultures were treated for 1 h with 50 µM ryanodine before the addition of 10 nM E2 for an additional 24 h. The hormone concentration used to study the neuritogenic effect of E2 was chosen based on previous studies of our laboratory (Gorosito et al., 2008; Scerbo et al., 2014).

After 2 DIV, neuronal cultures were fixed for 20 min with warm 4% paraformaldehyde in PBS containing 0.12 M sucrose and rinsed in PBS. Neurons were immunocytochemically stained with antibodies against β-tubulin class III (SDL.3D10). The details of the immunocytochemical procedure were as specified by Díaz et al. (1992). Briefly, the fixed cells were permeabilized in 0.2% Triton X-100 for 5 min at RT, preincubated with 5% BSA, incubated in mouse anti-β-tubulin class III (Sigma-Aldrich, St. Louis, MO, USA), rinsed in PBS, and finally incubated with appropriate biotinylated secondary antibody. Incubation with secondary antibody was followed by washing in PBS, incubation for 2 h in VECTASTAIN ABC immunoperoxidase reagent (Vector Laboratories, Burlingame, CA, USA), and a final reaction with 1.4 mM 3,3'-diaminobenzidine in phosphate buffer with H2O2. Coverslips were then dehydrated with ethanol, cleared with xylene, and mounted on glass slides for morphometric analysis. No immunostaining was detected when primary antibodies were replaced by 5% BSA.

#### Morphometric Analysis

The morphometric analysis of stained neuronal cultures was performed on digitized video images using JAVA as an image processor (Jandel Inc., Richmond, CA, USA) controlled by a host computer. Images were acquired through an optic microscope (Carl Zeiss, Germany). Microscope slides were coded, and the person conducting the analysis was blind to the experimental group. All labeled cells that could be identified as one individual neuron were measured in random fields at 40× magnification. Neural processes were classified as minor processes or axons according to accepted morphological criteria (Dotti et al., 1988; Blanco et al., 1990; Díaz et al., 1992). Minor processes are two or three short neurites that emerge from the cell body; axons are much longer, unique, thin, and relatively uniform in diameter. Neurons were considered to have developed an axon

washing for 2 h, the cultures were treated with the inhibitors for 1 h and were then pulsed for 15 min with 17β-estradiol (E2) and harvested for Western blotting. Top: ratio of readings for pERK/ERK bands in arbitrary densitometric units. Bottom: examples of immunoblots showing a decrease of hormone-induced ERK phosphorylation in cultures pretreated with the inhibitors. Molecular masses (kDa) for ERK1 and ERK2 are 44 and 42, respectively. Blots shown are representative of the mean ± SEM of 3–4 different cultures. (A) Nifedipine: ANOVA F(2,10) = 52.78; p ≤ 0.001. Least significant differences (LSDs) test indicated ∗∗∗p < 0.001 vs. control and p < 0.01 vs. E2. (B) Ryanodine: ANOVA F(2,6) = 5.856; p = 0.04. LSDs test indicated ∗p = 0.05 vs. control and p = 0.05 vs. E2. (C) 2-APB: ANOVA F(2,6) = 27.203; p ≤ 0.001. LSDs test indicated ∗∗∗p < 0.001 and ∗p = 0.05 vs. control and p = 0.01 vs. E2. (D) U-73122: ANOVA F(2,6) = 34.891; p ≤ 0.001. LSDs test indicated ∗∗∗p < 0.001 and ∗∗p = 0.01 vs. control and p ≤ 0.05 vs. E2.

if they showed one neurite three to five times longer than the rest (stage III of development). Soma area, length of minor processes, total axonal length, and the number of neurites per cell were recorded. At least 60 neurons were measured for every experimental condition in each culture; at least three separate cultures were made for every condition.

#### Statistical Analysis

Data were statistically evaluated by one-way ANOVA, followed by Fisher's Least Significant Difference (LSD) post hoc test (Statistica; StatSoft Inc., Tulsa, OK, USA) where p < 0.05 was considered statistically significant.

#### RESULTS

#### E2-Induced ERK1/2 Activation Is Mainly Mediated by RyRs

Our previous results suggested that E2-induced ERK1/2 phosphorylation is Ca2+-dependent. Here, we further

representative experiment out of six independent experiments.

investigated the Ca2+ response involved in ERK activation mediated by E2. Hypothalamic cultures grown with E2 for 48 h were washed in BM for 2 h and pre-treated with specific compounds for 1 h before a pulse of E2 for 15 min. In agreement with previous reports, E2 induced a strong phosphorylation of ERK at 15 min (**Figure 1**). This effect was completely abolished by inhibitory ryanodine (**Figure 1B**) and partially attenuated by nifedipine (**Figure 1A**), IP3R inhibitor 2-APB (**Figure 1C**), and a PLC inhibitor U-73122 (**Figure 1D**).

## E2 Induces Rapid Ca2+ Increase Depending on Ca2+ Influx and RyRs

As E2-activation of the ERK1/2 signaling cascade depends on extracellular as well as intracellular Ca2+ stores, we decided to characterize the Ca2+ signal generated by the hormone. The addition of agonist to cell cultures, loaded with the indicator Cal-520 AM and imaged in a Ca2+-containing buffer, induced fluorescence changes that were observed in neuronal soma as well as in minor processes (**Figure 2**). Seventeen out of

76 neurons (22.4%) from six independent experiments imaged in Ca2+-HBSS responded to E2 pulses within 15–100 s of treatment (average = 46.8 ± 6.60 s). These Ca2+ events often occur in repetitive oscillations, which display interspike intervals of 21.58 ± 8.83 s (9 out of 17 neurons). The amplitude in terms of ∆F/Fo was 0.154 ± 0.01. The total amount of Ca2+ mobilized, measured as the integrated area under the curve (AUC) for all E2-generated peaks, was 4.69 ± 1.03, which was 90.6% with respect to the control [remained endoplasmic reticulum Ca2+ content released by tg, a sarco/endoplasmic reticulum Ca2+- ATPase (SERCA) inhibitor].

This E2-induced Ca2+ signal was not observed in the absence of extracellular Ca2+. Moreover, we found that pre-incubation of neuronal cultures with inhibitory ryanodine suppressed E2-evoked Ca2+ release (**Figure 3A**). Importantly, under these conditions, the tg was able to mobilize amounts of Ca2+, measured as AUC, of 14.69 ± 3.45 (EGTA-containing buffer) and 35.05 ± 11.54 (ryanodine pre-incubation), five and nine times greater, respectively, than the amounts of Ca2+ mobilized by tg in E2-induced control neurons (**Figure 3B**). Moreover, nifedipine reduced the E2-induced Ca2+ increase more than 50% (∆F/Fo = 0.065 ± 0.011; and AUC = 10.779 ± 1.159, n = 4), which strongly suggests the participation of L-VGCCs in this signal (**Figures 3A,B**). Representative Ca2+ traces plotted as ∆F/Fo vs. time for EGTA, ryanodine, and nifedipine conditions are provided in **Supplementary Figure S1**.

Taken together, these results indicate that both Ca2+ influx and mobilization from intracellular stores contribute to the response.

#### E2-Induced Axonal Growth Depends on Ca2+ Signal Generated by RyRs

Finally, we tested whether the RyR-induced Ca2+ response is part of the signaling cascade that mediates the axogenic effect of E2. Neurons grew under the following conditions: in the presence and absence of E2 and pretreated with inhibitory ryanodine. After these treatments, the cells were grown for an additional 24 h period with (E2) or without E2 (control). In agreement with previous results, the morphometric analysis indicated that neurons grown under hormonal treatment show longer axons than neurons in control conditions without E2 (**Figure 4A**). Remarkably, blocking RyRs with ryanodine completely inhibited the E2-induced axogenesis (**Figure 4B**). Moreover, no significant differences were observed in the number of primary neurites, length of minor processes, or soma area per neuron resulting from E2 or ryanodine treatment (**Table 1**), confirming that the hormonal effect is restricted to axonal growth (Díaz et al., 1992; Cambiasso et al., 1995, 2000; Brito et al., 2004).

#### DISCUSSION

In the present study, results from ERK1/2 phosphorylation, Ca2+ imaging and neuronal growth consistently pointed to RyRs as the Ca2+ channels necessary to mediate activation of the MAPK/ERK pathway and the final axogenic effect induced by E2. Inhibitory ryanodine completely blocked E2-mediated Ca2+ transients, ERK1/2 phosphorylation and axonal outgrowth, which provides valuable evidence to propose that E2 mobilizes endoplasmic reticulum stores of Ca2+ through RyRs to activate the signaling cascades that finally affect the axonal elongation of hypothalamic neurons.

In regard to the impact of functional RyRs on cellular response, Dr Hidalgo's group showed that BDNF-induced neural plasticity requires functional RyRs activated by the Ca2+ induced Ca2+ release (CICR) mechanism to evoke the larger Ca2+ signaling needed to maintain changes during long-term memory storage (Adasme et al., 2011). CICR is a positive

(control) 10 nM 17β-estradiol in combination or not with 50 µM ryanodine (Ry) for the last 24 h of incubation (arrows indicate the axons of some neurons). (B) Mean of axonal length for each condition in (A). ANOVA F(3,12) = 4.51; p = 0.02. LSDs test indicated ∗p = 0.05 vs. control and p = 0.01 vs. E2. Data represent the mean ± SEM; n = 4 independent cultures. Scale bar: 100 µm.

TABLE 1 | Number of primary neurites, soma area, and length of minor processes of male hypothalamic neurons grown with or without 17β-estradiol (E2) in 3 combination or not with ryanodine.

Data represent the mean ± SEM; n = 4 independent cultures.

feedback mechanism by which cytoplasmic Ca2+ stimulates Ca2+ release from the endoplasmic reticulum through RyRs or IP3Rs (Bezprozvanny et al., 1991; Berridge et al., 2003; Seo et al., 2015).

Moreover, several studies have indicated a regulatory role for estrogens on RyRs activity, for instance, in the human eccrine sweat gland cell line NCL-SG3 (Muchekehu and Harvey, 2008), Cabrera Zapata et al. Estradiol-Induced Ca2+ Signaling and Axogenesis

ventricular myocytes (Yan et al., 2011), detrusor smooth muscle cells (Hristov et al., 2017), and dorsal root ganglion neurons (Ferrari et al., 2016; Khomula et al., 2017). Interestingly, Zhao X. et al. (2005) proposed, in a neuroblastoma cell line, an E2-mediated mechanism starting at the plasma membrane, by which rapid Ca2+ signaling potentiates the transcription of genes normally regulated by estrogens; RyRs, IP3Rs, and N-VGCCs, but not L-VGCCs, were involved in the process. Our results clearly show that RyRs are indispensable Ca2+ channels involved in the non-classical signaling events produced by E2 to generate axonal growth in hypothalamic neurons. However, RyRs also require a previous small increase in cytosolic Ca2+ from resting levels to activate and release Ca2+ by CICR (Hidalgo et al., 2005; Lanner et al., 2010). Since we found that removing extracellular Ca2+ or blocking the membrane channels L-VGCCs prevented the Ca2+ signaling induced by E2 and that inhibiting L-VGCCs also reduced ERK1/2 phosphorylation modulated by the steroid, we postulate that E2 initially induces a Ca2+ influx in hypothalamic neurons via L-VGCCs that then enables RyRs opening to generate the final and complete Ca2+ signaling event (Hidalgo, 2005; Calin-Jageman and Lee, 2008). Besides L-VGCCs, our results show that the PLC/IP3Rs system is involved in E2-induced ERK1/2 activation, since both 2-APB and U-73122 used as blockers of IP3Rs and PLC, respectively, produced a significant reduction in phosphorylation levels of the kinases in the presence of the hormone. The activation of PLC and Ca2+ release via IP3Rs induced by estrogens has been previously reported in different cellular systems (Chaban et al., 2004; Fricke et al., 2007).

L-VGCCs are the major route of Ca2+ entry into neurons and the most profusely studied and best characterized VGCC type by far, as they play a predominant role in the brain (Striessnig et al., 2014; Vega-Vela et al., 2017). Several studies report that E2 is able to modulate L-VGCCs activity (Bulayeva et al., 2005; Sarkar et al., 2008; Farkas et al., 2012; Feng et al., 2013). Wu et al. (2005) and Zhao L. et al. (2005) indicated that E2 induced rapid Ca2+ influx through L-VGCCs, which was required to activate the Src/ERK/CREB/Bcl-2 signaling pathway and finally mediated neuroprotective and neurotrophic responses in rat hippocampal and cortical neurons. The generation of this intracellular Ca2+ increase and the downstream activation of ERK depend on the presence of ERs in the membrane of rat hippocampal neurons (Wu et al., 2011). These membrane ER-expressing neurons represented 29% of the cultured cells and all of them co-expressed L-VGCCs. Consistently, our data from Ca2+ imaging experiments indicated that approximately 22% of hypothalamic neurons responded to E2.

Although previously it has been reported that E2 induces L-VGCCs-mediated Ca2+ influx, the question about how the hormone activates L-VGCCs remains open. Sarkar et al. (2008) reported that E2 potentiated the activity of L-VGCCs by directly binding to specific sites in the channel, independently of ERs. On the other hand, PI3K signaling cascade has been proposed as a candidate to link membrane ER activation with L-VGCCs aperture (Simoncini et al., 2000; Quignard et al., 2001; Wu et al., 2005), although our preceding work blocking PI3K with LY-294,002 did not prevent the axogenic effect of E2 in hypothalamic neurons (Gorosito and Cambiasso, 2008). It is important to note that we have previously reported the expression of ERα on the cell-surface of embryonic hypothalamic neurons (Gorosito et al., 2008), and that the membrane-impermeable E2-albumin construct (E2-BSA) was as effective as free E2 to generate ERK1/2 phosphorylation (Gorosito and Cambiasso, 2008) and axonal elongation (Cambiasso and Carrer, 2001), evidence that altogether indicate these processes respond to a membraneinitiated ERα-mediated mechanism.

Another question that arises is what other elements lead from the RyRs-mediated Ca2+ release to ERK1/2 activation. In Gorosito and Cambiasso (2008) we reported that PKC but not PKA nor CaMKII is required in the E2-induced MAPK-ERK pathway activation, since an inhibitor with specificity for the PKC Ca2+-dependent α and βI isoforms, Ro 32-0432, attenuated E2-modulated ERK1/2 phosphorylation and prevented the axogenic effect of the hormone. PKC activation by E2 has been found in different cell types, including breast cancer cells, hepatocytes, and cortical and hypothalamic neurons (Marino et al., 1998; Boyan et al., 2003; Cordey et al., 2003; Qiu et al., 2003). Ca2+-dependent PKC activation can then induce ERK1/2 phosphorylation via Src/Ras signaling (Cullen and Lockyer, 2002; Brandt et al., 2003; Roskoski, 2005).

Our results show that E2-induced cytosolic Ca2+ often increases as repetitive oscillations. This is in agreement with previous work demonstrating E2-induced intracellular Ca2+ oscillations that involved internal stores and PKA and PLC activity in neurons of the arcuate nucleus (Fricke et al., 2007). It is well known that the frequency of Ca2+ oscillation may depend on Ca2+ influx into the cell (Sneyd et al., 2004), SERCA activity (Falcke et al., 2003) and oscillating cytoplasmic IP3 concentration (Sneyd et al., 2006). Our data suggest that the frequency of Ca2+ oscillations in hypothalamic neurons stimulated by E2 mainly depends on Ca2+ entry. Leaving aside its modulation, our results allow us to speculate that the characteristic frequency encodes information to regulate the cellular response (axonal growth) mediated by the hormone (Dolmetsch et al., 1998).

In conclusion, we have provided new insights into the non-classical mechanisms triggered by estrogens and its axogenic effect in male rat hypothalamic neurons. The hormone induces ERK1/2 activation in a Ca2+-dependent manner. RyRs inhibition abolished this activation as well as axonal growth. The oscillatory Ca2+ signal generated by E2 required functional RyRs and L-VGCCs. This early Ca2+ response that underlies E2-induced RyRs and MAPK/ERK activation may transmit a finely tuned message into a neuronal development program, reflecting the need for tight control of a critical event during sexual differentiation of the male brain. The conjunction of XY genotype with adequate estrogen exposure levels at the time of hypothalamic neuronal differentiation may induce the growth of axons towards their appropriate targets. A complete and detailed understanding of the intracellular signaling mechanisms and neuronal processes mediated by estrogens will allow to improve current estrogen-based therapies, such as hormone replacement therapy in postmenopausal women, as well as to develop novel treatments to prevent and/or alleviate neurological pathologies based on its widely proven neuritogenic and neuroprotective effects.

#### AUTHOR CONTRIBUTIONS

LCZ, MB and MC made the conception and design of research, interpreted results of experiments, wrote the manuscript. LCZ performed all experiments and analyzed the data. LCZ and MB prepared the figures. MB and MC edited and revised the manuscript. MC drafted the manuscript.

#### FUNDING

This study was supported by grants from Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET;

#### REFERENCES

PIP 2013-2015), Agencia Nacional de Promoción Científica y Tecnológica (ANPCyT; PICT 2015 N◦ 1333) and Secretaria de Ciencia y Tecnología (SECyT)-Universidad Nacional de Córdoba (2016-2017), Argentina to MC and from CONICET (PIP 2014-2016) to MB.

#### ACKNOWLEDGMENTS

We thank for the technical support from INIMEC animal facility and Dr Quassollo for his invaluable help with imaging. Calcium imaging was generated at National Center of Microscopy, Universidad Nacional de Córdoba, Córdoba, Argentina.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fncel. 2019.00122/full#supplementary-material

receptor-negative HCC38 cells is membrane-mediated and inhibited by tamoxifen. Endocrinology 144, 1812–1824. doi: 10.1210/en.2002- 221018

fluxes. Proc. Natl. Acad. Sci. U S A 101, 1392–1396. doi: 10.1073/pnas.<PHONE_NUMBER>

**Conflict of Interest Statement**: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Cabrera Zapata, Bollo and Cambiasso. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Oxidative Stress in Retinal Degeneration Promoted by Constant LED Light

#### Maria M. Benedetto1,2 and Maria A. Contin1,2 \*

1Departamento de Química Biológica Ranwel Caputto, Facultad de Ciencias Químicas, Universidad Nacional de Córdoba, Córdoba, Argentina, 2Centro de Investigaciones en Química Biológica de Córdoba (CIQUIBIC), CONICET, Universidad Nacional de Córdoba, Córdoba, Argentina

Light pollution by artificial light, might accelerate retinal diseases and circadian asynchrony. The excess of light exposure is a growing problem in societies, so studies on the consequences of long-term exposure to low levels of light are needed to determine the effects on vision. The possibility to understand the molecular mechanisms of light damage will contribute to the knowledge about visual disorders related to defects in the phototransduction. Several animal models have been used to study retinal degeneration (RD) by light; however, some important aspects remain to be established. Previously, we demonstrated that cool white treatment of 200 lux light-emitting diode (LED) induces retinal transformation with rods and cones cell death and significant changes in opsin expression in the inner nuclear layer (INL) and ganglion cell layer (GCL). Therefore, to further develop describing the molecular pathways of RD, we have examined here the oxidative stress and the fatty acid composition in rat retinas maintained at constant light. We demonstrated the existence of oxidative reactions after 5 days in outer nuclear layer (ONL), corresponding to classical photoreceptors; catalase (CAT) enzyme activity did not show significant differences in all times studied and the fatty acid study showed that docosahexaenoic acid decreased after 4 days. Remarkably, the docosahexaenoic acid diminution showed a correlation with the rise in stearic acid indicating a possible association between them. We assumed that the reduction in docosahexaenoic acid may be affected by the oxidative stress in photoreceptors outer segment which in turn affects the stearic acid composition with consequences in the membrane properties. All these miss-regulation affects the photoreceptor survival through unknown mechanisms involved. We consider that oxidative stress might be one of the pathways implicated in RD promoted by light.

Keywords: retinal light damage, LED light, oxidative stress, fatty acid, electroretinogram

#### INTRODUCTION

The disturbance between the amount of reactive oxygen species (ROS) and antioxidants production is defined as oxidative stress. This imbalance produces tissue injury (Halliwell, 1994). The retina carries out the capture of light photons, and for this, it is exposed to suffer oxidative stress. However, it has many mechanisms to counteract these processes through the action of antioxidant enzymes as superoxide dismutase (SOD), catalase (CAT) and glutathione peroxidase (GSHPx) and vitamins as ascorbic acid, vitamin E, melanin (Iusifov et al., 1980; Scibior and Czeczot, 2006; Pavarino et al., 2013). The overexposure to light may

#### Edited by:

Gabriela Alejandra Salvador, Universidad Nacional del Sur, Argentina

#### Reviewed by:

Luis Enrique Politi, Instituto de Investigaciones Bioquímicas de Bahía Blanca (INIBIBB), Argentina Rocío Salceda, National Autonomous University of Mexico, Mexico

#### \*Correspondence:

Maria A. Contin<EMAIL_ADDRESS><EMAIL_ADDRESS>
Received: 07 December 2018 Accepted: 20 March 2019 Published: 11 April 2019

#### Citation:

Benedetto MM and Contin MA (2019) Oxidative Stress in Retinal Degeneration Promoted by Constant LED Light. Front. Cell. Neurosci. 13:139. doi: 10.3389/fncel.2019.00139 be one of the many factors that can induce the interruption of this homeostasis, promoting the injury of eye tissues, cell death or stimulating simultaneously antioxidant protection by up-regulation of antioxidant enzymes (Yusifov et al., 2000); however, when the equilibrium is broken the consequence is the induction of retinal degeneration (RD). In mammals, in normal light conditions, the retina fulfills two main roles: the vision over rods and cones activity and the non-image forming tasks including circadian entrainment, pupillary response to light, secretion of melatonin and sleep regulation (Berson et al., 2002; Rollag et al., 2003; Guido et al., 2010; Sikka et al., 2014). Light stimulates photoreceptor cells activating the phototransduction cascade which promotes hyperpolarization (Sung and Chuang, 2010). Light overexposure-induced RD might induce retinal task deficits with consequences in the secretion of melatonin, desynchronization of rhythms such as sleep/wake, among others. The use of different sources of artificial light is increasing in actual society but this promotes changes in human behavior. Furthermore, the use of technologies such as light-emitting diode (LED), smart TVs and cell-phones, promote an excess of blue light exposure especially at night. Albeit the effects of artificial illumination are unknown, it may have a strong impact on retinal functions with negative consequences on people's health. Considering that the excess of light exposure constitutes an upcoming polluter, it is clearly an emerging public health issue. Different animal models have been used by researchers in order to study the processes of RD promoted by light exposure.  In retina and retinal pigment epithelium (RPE) cells, the exposure to blue light inhibits the mitochondrial enzymes and cytochrome oxidase expression inducing retinal damage (Chen, 1993; Cai et al., 2000; Roehlecke et al., 2011) suggesting the existence of oxidative stress mechanisms as part of the retinal cell death mechanism induced by light exposure. Recently, Nakamura et al. (2017) established that LED light exposure during 2 h (800 lx) induced retinal damage where oxidative stress was partially involved. However, the consequences of constant low light exposure in which the phototransduction mechanism is constantly activated for long time remain unknown. In this regard, we have established and characterized a model of RD induced by a constant exposure to LED. We have showed that classical photoreceptor cell death without caspase-3 activation and a gradual increase in levels of rhodopsin-phospho-Ser334 as a result of light exposure, suggesting that constant light produces changes in the regulation of phototransduction in rods (Contín et al., 2013).

However, retinal ganglion cells (RGCs) do not die after constant light treatment (LL), suggesting a neuroprotection mechanism also involved in our model (Benedetto et al., 2017). So, the key questions in the present work are: does the overexposure at constant low levels of LED light yield oxidative stress? What is the kinetic of oxidative stress production if the exposure is constant? Therefore, the goals of this work were to investigate the existence of oxidative stress, fatty acid composition and retinal function during the different days of LL stimulation in retinas of Wistar rats.

### MATERIALS AND METHODS

#### Animals

All animal procedures were performed in accordance with the protocol approved by the local animal committee (School of Chemistry, UNC, Exp. 2018-740), in accordance with the ARVO statement for the use of animals in ophthalmic and vision research. Wistar rats from 12 to 15 weeks of age were maintained with food and water ad libitum and illumination cycle from 12:12 h (light/dark) with white fluorescent light on (∼50 lux) from Zeitgeber time (ZT) 0–12 from the time they were born, up to the experiment.

### Light Damage

#### Constant Light

RD was induced as described by Contín et al. (2013). Briefly, rats were exposed to constant light in boxes with LED lamps (EVERLIGHT Electronic Co., Ltd. T-13/4 3294-15/T2C9-1HMB, color temperature of 5,500 K) in the inner upper surface and temperature-controlled at 24 ± 1 ◦C. At the level of the rats' eyes, 200 lx were measured with a light meter (model 401036; Extech Instruments Corp., Waltham, MA, USA). After 1 to 8 days of constant light stimulation (LL1–LL8) the animals were killed in a CO2 chamber at ZT6. Controls in light dark cycle (LD), with LED or fluorescent light (RT) and constant darkness (DD) were exposed for 7 days.

#### Dark Period Protocol

Animals were subjected to a constant light during 8 days with periods of dark during 2, 4, 6, 10 and 12 h every day at the beginning of the subjective night (ZT6) under identical conditions as animals exposed to LL, in the temperaturecontrolled stress box at 24 ± 1 ◦C.

### Electroretinograms (ERGs)

The methods employed for Scotopic electroretinogram (ERG) were as previously described by Dorfman et al. (2014) using an ERG machine (Akonic BIOPC, Buenos Aires, Argentina). Briefly, first animals were adapted to dark for 20 min. Then, they were anesthetized with an intraperitoneal injection containing a solution of xylazine hydrochloride (2 mg/kg) and ketamine hydrochloride (150 mg/kg). Pupils were dilated with tropicamide (1% Alcon Laboratories) and, in order to prevent eye dehydration and permit electrical contact activity when the electrode is recording, the cornea was irrigated with proparacaine hydrochloride (0.5% Alcon Laboratories). Both eyes were recorded simultaneously applying flashes of white light (5 ms, 0.1 Hz) from a photostimulator set at maximum brightness (3 cd s/m2 without filter). Then, the recordings were amplified and filtered (1.5 Hz low-pass filter, 300 Hz high-pass filter, notch filter activated). An average of 10 responses for each eye was measured. Mean ''a'' and ''b'' waves peak latencies and amplitudes of the responses from each group of rats were compared.

#### Outer Nuclear Layer Analysis

The retinal fixation method, sectioning and nuclear quantification were as previously described (Contín et al., 2013). Briefly, rats' eyes were fixed overnight at 4◦C in 4% (W/V) paraformaldehyde in 100 mM sodium phosphate buffer (PBS, pH 7.3). Then, they were cryoprotected in sucrose and mounted in an optimal cutting temperature compound (OCT; Tissue-Tekr Sakura). Retinal sections were cut along the horizontal meridian (nasal-temporal). The sections were stained with 1% Hoechst (33258 Sigma Aldrich) for 5 min and photographed using a confocal microscope (Olympus FV1200, Japan) at 40× magnification. The nuclei were counted in ''left, middle left, middle right and right'' designated areas from five different animals per treatment using the software ImageJ (v. 1.45) and the plugin ''Automatic Nuclei Counter''.

#### Superoxide Production

Dihydroethidium [(DHE; sigma 37291-25 mg dihydroethidium at 10-mg/mL stock solution in dimethylsulfoxide)], a redoxsensitive probe, was used to detect superoxide generation as previously described (Peng et al., 2011). Briefly, eyes were removed, washed in PBS solution and then incubated with 1 mM DHE for 12 h at room temperature in PBS. Then the eyes were incubated in paraformaldehyde 4% for 12 h at room temperature, harvested and quikly frozen in liquid nitrogen for cryosection (Leica CM 1950, Leica Microsystems Ltd, Wetzlar, Germany). The cryosections (10 µm) were analyzed by confocal microscopy (Olympus FV300, Japan). The DHE molecules enter the cells and are oxidized by superoxide contained inside cells, to ethidium (Et) which is fluorescent. This product is retained in the cell allowing the estimation of cellular superoxide production.

#### Oxidative Stress Quantification

ROS molecules were determined by flow cytometry with 2 0 ,70 -dichlorofluorescein diacetate (DCFH-DA) probe (D6883 Sigma). DCFH-DA is sensitive to oxidation and is nonfluorescent, it may be oxidized by ROS and peroxides and become a fluorescent molecule, DCFH (Gomes et al., 2005). Briefly, retinas dissected and dissociated in 0.25% (W/V) trypsin (Life Technology, Inc., Carlsbad, CA, USA) for 7 min at 37◦C were centrifuged at 3,000 rpm for 3 min, incubated with 200 µl of trypsin inhibitor (STI) for 3 min and washed twice with PBS 1×. Then, they were suspended in 200 µl of PBS 1×, incubated with DCFH-DA (5 µM) for 60 min at 37◦C in dark, washed twice with PBS 1× and analyzed on a Becton-Dickinson FACS flow cytometer; the excitation and emission wavelengths were set at 488 and 525 nm, respectively. Data were analyzed with FlowJo software (LC, Ashland, Ore). Three independent experiments were performed and the results were expressed as mean ± standard error (SE) in arbitrary units of DCFH fluorescence intensity.

### Catalase Activity Detection

Catalase (CAT) activity was assessed according to Aebi (1984). Briefly, individual retinas were homogenized in PBS 1×. Then, 1.3 ml of Buffer Phosphate (50 mM, pH 7.4) and 1.25 ml of distilled H2O were added to 300 µl of previously homogenized sample. Subsequently, 150 µl of H2O2 (300 mM) was added and the spectrophotometric changes were detected by measuring the absorbance at 240 nm for 1 min. The protein concentration was determined by Bradford (1976).

### Lipid Extraction From Rat Retinas

Lipids were extracted from the samples as Folch' method slightly modified (Folch et al., 1957). Briefly, collected retinas were homogenized in 200 µl of MiliQ H2O and 3 ml of a combination of Chloroform: methanol [(2:1; v/v)] and 600 µl of MiliQ H2O were added and vortexed vigorously for 5 min and centrifuged at 2,000 rpm at 15◦C for 10 min. The layer containing chloroform was collected and 1.5 ml of a mixture of chloroform: methanol: H2O [(3:48:47; v/v/v)] was added, mixed gently and centrifuged 10 min at 2,000 rpm. Finally, chloroform was dried under a stream of N2 and the extracts stored for further experiments.

#### Fatty Acid Methyl Esters Analysis by Gas Chromatography/Mass Spectrometry With Ion-Trap Detector

Extracted lipids were transmethylated to fatty acid methyl esters (FAMEs) with sodium methoxide and toluene [(2:1; v/v)] overnight at 4◦C. FAMEs were then extracted with hexane and analyzed by using a GC/MS (Saturnr 2000 GC/MS, Agilent, Santa Clara, USA) equipped with an ion-trap detector and a fused capillary column (HP-5MS, 30 mm × 0.25 mm i.d × 0.25 µm film thickness, Agilent, Santa Clara, USA). The carrier gas used was Helium (flow rate 1.0 ml/min); the detector and injector temperature were maintained at 150◦C and 300◦C, respectively. The injection (split-splitless injector) volume of the sample was 1 µl with a split ratio of 1:50. The oven was programmed as follows: 170◦C for 3 min—200◦C at the rate of 1.2◦C/min—10◦C/min to 240◦C—80◦C/min to 280◦C and 80◦C/min to 300◦C with a final hold for 46.7 min.

The column temperature was programmed as follows: 170◦C; 1.2◦ /min to 200◦C—10◦ /min to 240◦C—80◦ /min to 2,800◦C—80◦ /min to 300◦C and hold at 300◦C for 4 min. Then, FAMEs were identified comparing mass spectra with dates in the library (National Institute of Standards and Technology, Gaithersburg, MD, USA). Furthermore, we compared FAMEs retention times with the commercial standard Supelco 37-Component FAME Mix (Sigma Aldrich, St. Louis, MO, USA).

#### Statistical Analysis

Statistical analysis was carried out using the Infostat software (Version 2017, InfoStat Group, FCA, National University of Córdoba, Argentina). One-way analysis of variance (ANOVA) was used for statistical comparisons. In the text and figure, data are expressed as mean ± SD or SE. It was considered

statistically significant a p-value < 0.05. The normality and homogeneity of the variance assumptions were proved with Shapiro–Wilks and Levene tests, respectively. Duncan post hoc test with a p value < 0.05 were considered statistically significant. A non-parametric Kruskal-Wallis test was performed when the data did not comply with the assumptions of the ANOVA. Spearman correlation coefficients were calculated to determine associations between the content of fatty acids.

#### RESULTS

#### Structural Retinal Analysis

In order to know the kinetic of cell death events we analyzed the retinal layers, oxidative stress, CAT activity, fatty acid composition and ERG activity in animals exposed to constant light. The analysis of retinal structure revealed a reduction in outer nuclear layer (ONL) thickness at LL6 and LL8 compared with animals reared in DD (**Figures 1A–C**). The quantification of nuclei in this area showed a significant reduction after 6 and 8 days of constant light (583.04 ± 45.31 and 580.10 ± 45.31 nuclei, respectively) compared with DD and RT [(1052.19 ± 38.41 and 1007.63 ± 54.32 nuclei, respectively, p < 0.005, **Figure 1D**)]; indicating that between LL0 to LL6 are being carried out the photoreceptor cell death pathways.

### Oxidative Stress Study

#### DHE Determination

In order to know if retinas from animals exposed to constant light have oxidative stress, DHE was measured in retinal sections as indicated in ''Materials and Methods'' section. DHE and superoxide anion reaction produces ethidium (E), which binds to DNA rising the fluorescence in the cells (Gomes et al., 2005; Fernandes et al., 2006). Here, we show that retinas from animals exposed for 2 days of LL (LL2) have fluorescent label in few cells in ONL; however, at 8 days of LL (LL8) a significant increment of DHE label was observed (**Figures 2A–C**), correlating with ONL cell reduction (**Figure 1**). Besides the clear staining in ONL, no positive labeled in other retinal layer were observed in any time of LL studied, indicating the existence of oxidized products specifically in this layer.

#### Reactive Oxygen Species Quantification

To determine the kinetic of oxidative stress events occurred in ONL of retinas exposed to LL, ROS production was determined

by flow cytometry using the DCFH-DA probe as indicated in ''Materials and Methods'' section. At 2 and 4 days of LL no significant increases in fluorescence were observed (124.70 ± 44.94 and 160.69 ± 16.9%, respectively). However, constant light treatment increases the levels of fluorescence, showing significant changes after 5 days of LL (LL5) where the production of ROS was maximal (372.16 ± 113.44%) respect to control in DD (100%). Although at 7 and 8 days of LL the fluorescence production decreased in correlation with cell death in the ONL (287.97 ± 221.55 and 211.00 ± 80.80%, respectively), ROS production was greater than in DD (**Figure 2D**).

#### Catalase Activity Determination

CAT enzyme is known to contribute to H2O2 detoxification in the retina and the inhibition of its activity increases H2O2 concentration 2.5-fold, which cannot make up for the GPX activity (Ohta et al., 1996) the enzyme being an antioxidant factor in retina. In order to investigate the CAT activity and its association in the prevention of oxidative stress in constant light exposed rats, it was assessed as indicated in ''Materials and Methods'' section. Although constant light stimuli increased lightly CAT activity at 6 and 8 days of LL (7.22 ± 3.95 and 5.96 ± 3.27, respectively) with respect to LL 2 and 4 (3.38 ± 1.44 and 4.35 ± 2.24, respectively), reaching the levels of DD and RT (5.82 ± 2.55 and 6.96 ± 4.30, respectively) there were no significant differences in any times of LL measured (**Figure 3**), suggesting that CAT is affected by the desynchronization of animals by constant light exposure.

#### Retinal Fatty Acid

The outer segment membrane of retinal rods and cones contain high polyunsaturated fatty acids (PUFAs) where docosahexaenoic (22–6 n-3, DHA) and arachidonic acid (20:4 n-6, AA) are the major species present. Because PUFAs are a target for oxidation, the analysis by GC-MS in rat retinas from animal exposure to different LL allow us to determine if there are changes in the membrane components during RD as a result of light exposure. Individual fatty acids were identified comparing the mass spectra with those present in the NIST library and their retention time with a commercial standard (see ''Materials and Methods''). Representative GC/MS analyses of fatty acids from retinas of control animals and exposed to light (LL) are shown in **Figure 4A**. Palmitic (16:0, PA) and arachidonic acid (20:4 n-6, AA), did not show statistical differences in any times of LL studied (**Figures 4B,D**). However, stearic acid (18:0, EA) shows increased levels with respect to control at LL4, maintaining this higher level at LL6 and LL8 (**Figure 4C**) in association with the DHA decrease which was significant at LL 4 and 8

(**Figure 4E**). In order to determine if the variations were related, we perform a correlation analysis between EA and DHA in control animals (DD) and animals exposed to 4, 6 and 8 days to LL. **Figures 5A–D** show no correlation in control animals maintained in dark (DD) and in LL6, however, there is a positive correlation in LL4 (ρ = 1) and negative correlation in LL8 (ρ = −0.8).

#### Electroretinograms Responses

Previously we demonstrated retinal functional alterations in the scotopic ERG responses, showing that throughout the days of LL, the two principal waves ''a'' and ''b'' tend to decrease their amplitudes and increase their latency time, reaching abolished records after LL4, indicating that photoreceptor cells fail between the first 4 days of LL (Quintana et al., 2016). Here, to study the effects of light interruption by darkness, the animals were exposed to constant light with 2, 4, 6, 10 or 12 h of dark during the subjective night as indicated in ''Materials and Methods'' section. As shown in **Figure 5**, with 2, 4 and 6 h of rest in darkness per day during subjective night, the ERG show abolished records as we described before, however, at 8 and 10 h of dark the ERG began to restore, reaching normal values at 12 h. This result indicates that including hours of rest in darkness during the subjective night the effects of light exposure may be less harmful than constant stimuli.

#### DISCUSSION

The excess of artificial light exposure constitutes a problem that is emergent worldwide. The behavior of excessive exposure to light might have effects on the vision, promoting RD and circadian asynchrony in healthy population. We have previously characterized a RD model that may provide the possibility to study specific events associated with RD under conditions of low light with LED sources, at constant or long-term periods of exposure (LP context). We demonstrated that photoreceptor cells died along the days of continuous light stimuli through a caspase-3 independent mechanism. Rhodopsin analysis did not show a reduction in protein levels, however, rhodopsin-phospho-Ser334 increased gradually with the days of LL exposure indicating increasing levels in the phosphorylation. The study of rhodopsin allows us to investigate the role of phototransduction mechanism in the RD model suggesting that rhodopsin dysregulation could be involved in one of the ways of RD (Contín et al., 2013). Furthermore, in the inner retina, we demonstrated a relocation of non-visual photopigments (OPN4 and OPN5); however, inner retinal cells survival were not affected, indicating a compensatory mechanism of protection with possible changes in synchronization of circadian rhythms (Benedetto et al., 2017). To further describe the molecular pathways of cell death, in this work, we have examined the oxidative stress involvement in the effects of constant exposure to LED sources. Results from both DHE and DCFH-DA probe analysis demonstrate the existence of oxidative reactions in LL exposed rats (**Figure 2**). The ONL label of DHE indicates the occurrence of oxidative stress only in this layer, discarding stress mechanisms in other cells of the retina, coinciding with cell death study. DCFH-DA analysis shows that the production of ROS increases significantly after LL5 with higher levels at this day of LL (**Figure 2D**) indicating oxidative stress from 5 days of constant light. Powerful defenses against oxidative stress are the effect of Vitamin E, Ascorbic Acid, and melanin as well as the activity of antioxidant enzyme superoxide dismutase (SOD), CAT and GSHPx (Yusifov et al., 2000). Here, we demonstrated that CAT activity does not present significant differences in all times studied (**Figure 3**) showing a light diminution at LL2 to LL4 with respect to controls in DD and LD treatments; however, at LL6 and LL8 the CAT activity increases showing levels similar to the controls. Previous studies have demonstrated that CAT enzyme is finely regulated by the circadian rhythms; CAT bioinformatic and mRNA studies have been shown putative E-box sites in CAT and GPx regulatory regions; furthermore, CAT mRNA expression and enzyme activity shows circadian rhythmicity with higher levels at the end of the day (Navigatore Fonzo et al., 2009; Kharwar and Haldar, 2012; Lacoste et al., 2017). Sani et al. (2006) have demonstrated that CAT activity shows changes across a 24-h period in mouse. Thus, we hypothesize that constant light might produce a complex and multifactorial biological process inducing progressive desynchronization of circadian rhythm with concomitant antioxidant function altered; and therefore, the break of the equilibrium between ROS over-production and antioxidant processes. However, further studies are necessary in order to conclude an association between them.

The vertebrate retina contains high concentration of PUFA; particularly, in the outer segment membrane of retinal rod and cones, the PUFAs DHA and AA are the major species examined (Fliesler and Anderson, 1983; Giusto et al., 2000; SanGiovanni and Chew, 2005). PUFAs are essential for maintaining the appropriate fluidity of the membrane, necessary for efficient phototransduction cascade (Brown, 1994; Gawrisch and Soubias, 2008). The composition of membrane lipids and their direct interaction with proteins

play an important role in the modulation of the rhodopsin function (Salas-Estrada et al., 2018). A good retinal function is subject to adequate membrane structure which contains high levels of DHA. Anderson and Penn (2004) proposed, as neuroprotective adaptive responses, changes in DHA levels at different environmental conditions with the aim to control the number of photons captured by rhodopsin molecule. In the fatty acid n-3-deficient rod outer segment, it has been shown that reduced rhodopsin transducin (Gt) coupling, reduced cGMP phosphodiesterase activity, and slower formation of metarhodopsin II-Gt complex, relative to the animals fed with n-3-adequate diet, explained the reduced activity of rod phototransduction in these animals (Anderson and Penn, 2004). These findings support the so called ''photostasis'' which infers the morphological and biochemistry adaptation to capture a constant number of photons. The constant light exposure may

break the adaptive mechanism and promote several pathways of cell death.

DHA and AA are obtained from two ways: (a) the diet or (b) synthesized from alpha-linolenic (18:3n3) and linoleic acids (18:2n6). DHA is transported from the liver to the RPE cells (Scott and Bazan, 1989). In rat rods, DHA is essential for normal development of function (Benolken et al., 1973; Wheeler et al., 1975) where rat deficient in PUFA showed reduced ERG responses (Jeffrey et al., 2002). DHA located mainly in photoreceptors represents 50% of the total fatty acid and recently it has been demonstrated that the membrane of rod outer segment has higher levels of PUFA and Very Long Chain-PUFA (VLC-PUFA) in rod-dominant rats than conedominant, suggesting that rods and cones do not have equal lipid requirements (Agbaga et al., 2018). The cell membranes are susceptible to oxidation due to high level of unsaturation of PUFAs, furthermore, retina membranes exposed to light, elevate the concentration of oxygen with the presence of rhodopsin photo-bleached products that increment the risk (Kagan et al., 1981; Wiegand et al., 1983; Rózanowska and Sarna, 2005; Hunter et al., 2012). Earlier study revealed that lipid peroxidation reactions in light-induced RD induce the specific loss of DHA from rod outer segment membranes during constant illumination with an increase in the production of lipid hydroperoxides. None of the other fatty acids, including the AA, changed significantly over the 3-day time periods; concluding that light mediates the peroxidation of PUFAS in outer segment membrane acids and supporting the hypothesis that peroxidation is involved in retinal light degeneration (Wiegand et al., 1983). So, as we exposed the rat retinas to constant light at 200 lux, and we demonstrated increase in ROS significantly at LL5, we decided to study if fatty acid composition is altered belonging to the days in LL, specially by oxidative stress processes. Results from GC-MS analysis shows that PA (saturated) and AA (unsaturated) do not show statistical changes with the days of LL. However, DHA (unsaturated) decreased with the LL exposure with statistical significance at LL4, 6 and 8 compared with controls in LD and RT (**Figure 4E**). Remarkably, the decrease of DHA draws a parallel with an SA increase at the same day of LL (**Figure 4C**). The study of the correlation between both fatty acids shows an association between them in LL-treated retinas; instead, we did not find any association in control animals (DD). We assumed that the reduction in DHA may be affected by the oxidative stress in photoreceptor outer segment which in turn affects the photoreceptor survival through unknown different miss-regulation mechanism involved. The hepatic metabolism, through desaturation and elongation steps, influences the fatty acid composition (Engler et al., 2000). Diabetes, atherosclerosis, neurological disorders, cancer and others are enhanced by the disturbances in fatty acid content (see Zolfaghari and Ross, 2003). One fatty acid biosynthesis pathway begins with the desaturation of PA and SA; in this way, delta-9-desaturase (∆9-desaturase) is the key enzyme necessary for the conversion from palmitic to palmitoleic acid (16:1n-7) and stearic to oleic acid (18:1n-9). ∆6 and ∆5-desaturase enzymes are required for the metabolism of essential fatty acids, linoleic (18:2n-6) and alpha-linolenic (18:3n-3), to long-chain PUFAs (LCP-DHA). Hormonal, dietary factors, peroxisomal proliferators and developmental processes can affect the activity of ∆9-desaturase, which alters monounsaturated and PUFA composition (Ntambi, 1995, 1999). In hypertensive rats, DHAfed ∆9-desaturase activity decreases at 53% suggesting that dietary DHA influences properties and function of cellular membranes due to changes in fatty acid composition (Engler et al., 2000). Animals with ∆9-desaturase activity dysregulation show changes in the relation of stearic:oleic acid with effects over membrane fluidity and function (Ntambi, 1995, 1999). Lai et al. (2010) saw that patients with BCD (Bietti crystalline dystrophy; a retinal degenerative disease) have higher concentration of stearic acid and lower octadecadienoic acid (18:1n-9) than healthy animals. Furthermore, the activity of ∆9-desaturase and the concentration of monounsaturated fatty acids were lower in BCD animals suggesting abnormalities in lipid metabolism demonstrating a direct relation between rising levels of stearic acid, lowering fatty acid concentration and retinal dysfunction (Lai et al., 2010). We think that reduction of DHA levels by oxidation in LL rats may affect the desaturase activity in retina causing the rising of stearic acid accumulation which might affect the membrane proprieties giving more rigidity to the outer segment. It may explain part of the cell death mechanism induced by oxidative stress. Furthermore, the excess of light may alter the levels of Neuroprotectin D1 (NPD1) due to the DHA oxidation. NPD1 is a biologically active DHA derivative which induces downstream pro-survival pathways such as gene expression, pro-apoptotic gene suppression and pro-inflammatory responses, among others. The diminution of DHA may alter these ways across inducing photoreceptor cell death (see Asatryan and Bazan, 2017). Other studies are required to better understand the effects of DHA reduction; nevertheless, analyzing the time course of oxidative stress, we consider that it is not the only pathway involved, because the production of DHE and ROS have a maximum expression after LL5-6 (**Figure 2**), and the reduction of ONL occurs between LL6 and

LL8; however, the two principal waves of ERG, a and b, tend to decrease the amplitudes and increase the latency time during light stimuli, reaching abolished records at LL4 (Quintana et al., 2016). All these results suggest that a dysfunction of retinal electrical activity occurs previously to redox imbalance. It has been demonstrated that low intensities of light stimuli need the activation of photopigments and phototransduction to induce RD (Hao et al., 2002; Grimm and Remé, 2013), suggesting that impairment of the phototransduction mechanism could be responsible for cell death. Besides, previously we demonstrated the existence of more phosphorylated rhodopsin (rhodopsinphospho-Ser334) after LL2 with a higher level at LL7, supporting the idea that changes in phototransduction cascade are also involved (Contín et al., 2013). Here, we show scotopic ERG responses abolished until 6 h of rest darkness during subjective night; however, after 8 h of dark the activity begin to restore, reaching normal values at 12 h (**Figure 6**). These results suggest that the existence of regulatory mechanisms tend to revert or prevent the process of cell death when the retina is maintained in rest (notice the difference between 10 and 12 h of darkness); nevertheless, when light exposure is prolonged a threshold is exceeded promoting a chain of different cell death pathways. Further studies are necessary to determine the role of opsinmediated RD processes in this model; however, we think that retinal dysfunction during the 1st days of LL (LL1-LL4) is promoted by phototransduction processes which may induce other pathways of cell death as the oxidative stress occurr. Therefore, prevention therapies with antioxidants would not completely solve RD by LED light.

#### ETHICS STATEMENT

Comité Institucional de Cuidado y Uso de Animales de Laboratorio en el ambito de la Facultad de Ciencias ´ Químicas—CICUAL-FCQ-en los proyectos científicos -fs. 6-EXP-UNC:0007526/201 8 Number 740.

#### AUTHOR CONTRIBUTIONS

MB has contributed with the hypothesis, has performed the experiments, the analysis of the results and discussion. MC is the corresponding author, and had written the manuscript.

#### FUNDING

This work has been supported by Consejo Nacional de Investigaciones Científicas y Tecnológicas de la República Argentina (CONICET), Secretaria de Ciencia y

#### REFERENCES

Tecnología—Universidad Nacional de Córdoba (SeCyT-UNC) and by Agencia Nacional de Promoción Científica y Técnica (FONCyT, PICT 2012 No. 021).

#### ACKNOWLEDGMENTS

We are grateful to Dr Pablo Yunes, Dr Carlos R. Mas and Dr Cecilia Sampedro for technical support and Rosa Andrada for animal facility management and to Lucia Carlota Pedraza for doing the language quality check of this manuscript.

retinal damage model. Biol. Pharm. Bull. 40, 1219–1225. doi: 10.1248/bpb. b16-01008

**Conflict of Interest Statement**: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Benedetto and Contin. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Inhibition of SIRT2 by Targeting GSK3β-Mediated Phosphorylation Alleviates SIRT2 Toxicity in SH-SY5Y Cells

#### Shuhu Liu† , Zhihua Zhou† , Ling Zhang, Siying Meng, Shuji Li and Xuemin Wang\*

Key Laboratory of Mental Health of the Ministry of Education, Guangdong Province Key Laboratory of Psychiatric Disorders, Department of Neurobiology, School of Basic Medical Sciences, Southern Medical University, Guangzhou, China

Sirtuin 2 (SIRT2) is thought to be important in the pathogenesis of Parkinson's disease (PD), and the inhibition of SIRT2 rescues α-synuclein toxicity in a cellular model of PD. Recent studies have focused on identifying inhibitors of SIRT2, but little is known about the processes that directly regulate its function. GSK3β is a serine/threonine protein kinase that affects a wide range of biological functions, and it is localized in Lewy bodies (LBs). Therefore, we investigated whether SIRT2 is regulated by GSK3β and enhances cell death in PD. In the present study, Western blot showed that total SIRT2 levels did not change noticeably in a cellular model of PD but that SIRT2 phosphorylation was increased, and GSK3β activity was elevated. In addition, mass spectrometry (MS) studies indicated that SIRT2 was phosphorylated by GSK3β at three specific sites. Phospho- or dephospho-mimicking studies demonstrated that this postmodification (phosphorylation) increased SIRT2 toxicity in SH-SY5Y cells. Collectively, our findings identify a posttranslational mechanism that controls SIRT2 function in PD and provide evidence for a novel regulatory pathway involving GSK3β, SIRT2, and α-synuclein.

#### Edited by:

Gabriela Alejandra Salvador, Universidad Nacional del Sur, Argentina

#### Reviewed by:

Diego Albani, Istituto Di Ricerche Farmacologiche Mario Negri, Italy Ulkan Kilic, University of Health Sciences, Turkey

#### \*Correspondence:

Xuemin Wang<EMAIL_ADDRESS>
†These authors have contributed equally to this work

> Received: 06 October 2018 Accepted: 08 April 2019 Published: 24 April 2019

#### Citation:

Liu S, Zhou Z, Zhang L, Meng S, Li S and Wang X (2019) Inhibition of SIRT2 by Targeting GSK3β-Mediated Phosphorylation Alleviates SIRT2 Toxicity in SH-SY5Y Cells. Front. Cell. Neurosci. 13:148. doi: 10.3389/fncel.2019.00148 Keywords: SIRT2, GSK3β, 6-OHDA, PD, neuroprotection

## INTRODUCTION

Parkinson's disease (PD) is a long-term neurodegenerative disorder that mainly affects the motor system. Since it was first reported by James Parkinson in 1817 (Lees, 2007), PD has become the second most common neurodegenerative disorder (Dawson and Dawson, 2003). The disease is characterized by the loss of dopaminergic neurons in the substantia nigra, and the pathological hallmark is the accumulation of α-synuclein protein aggregates called Lewy bodies (LBs). It is believed that the number and size of α-synuclein aggregates affect the progression of PD. However, the exact molecular mechanisms that contribute to dopaminergic neuron loss remain to be clarified.

To explore the molecular mechanisms of PD, we focused on sirtuins, a family of class III nicotinamide adenine dinucleotide-dependent deacetylases that deacetylate histone and nonhistone proteins. SIRT2 is one of seven members of the sirtuin family, and it plays diverse roles in cellular metabolism and aging. SIRT2 is highly expressed in the central nervous system (Maxwell et al., 2011; Zhu et al., 2012) and is located in the cytoplasm (North et al., 2003), the nucleus (Dryden et al., 2003; North et al., 2003) and mitochondria (Liu et al., 2017). It has been reported that SIRT2 plays essential roles in PD, and SIRT2 inhibitors rescue α-synuclein-mediated toxicity (Outeiro et al., 2007; Chen et al., 2015). One SIRT2 inhibitor, AK7, even protects against MPTP neurotoxicity in mice (Chen et al., 2015). The protective effects of SIRT2 inhibition are mediated through a number of pathways. First, SIRT2 can directly deacetylate α-synuclein to exacerbate α-synuclein toxicity in vivo (de Oliveira et al., 2017). Second, SIRT2 inhibition achieves neuroprotection by reducing sterol levels via the decreased nuclear trafficking of SREBP-2 (Luthi-Carter et al., 2010). Third, SIRT2 inhibition may be neuroprotective in PD by modulating a redox network (Wang et al., 2015; Guan et al., 2016). Although SIRT2 plays a key role in the development of PD, we still do not know how SIRT2 itself is regulated during the development of this disease. It has been reported that SIRT2 is a phosphorylation substrate of CDK5, which modulates the activity of SIRT2 (Pandithage et al., 2008). However, there have been no reports that CDK5 can regulate the activity of SIRT2 in PD. To obtain further insight into the mechanism by which SIRT2 is regulated, we sought to identify novel upstream kinases of SIRT2. GSK3β and CDK5 are two kinases at the center of research on Alzheimer's disease, and they share the same substrate (Wen et al., 2008). Therefore, we hypothesized that SIRT2 may be a substrate of GSK3β.

GSK3β is a serine/threonine protein kinase that is activated by neurotoxins (Hongo et al., 2012; Hernandez-Baltazar et al., 2013; Zhao et al., 2016) and PD-associated gene mutations (Wang et al., 2013; Kawakami et al., 2014). Additionally, in the postmortem PD brain, GSK3β is localized in LBs, as is phosphorylated GSK3β (Ser9; Nagao and Hayashi, 2009). Furthermore, in a study of a group of 251 Spanish patients with PD, Infante et al. (2010) found that a GSK3β (rs6438552) TT genotype, which has been shown to produce a more active isoform (Kwok et al., 2005), is associated with an elevated risk of PD. Thus, GSK3β is important in the development of PD.  These results indicate that GSK3β is a critical mediator of 6-OHDA/MPP (+) induced neurotoxicity.

Based on the above information, we propose that SIRT2 may be phosphorylated by GSK3β during the development of PD. Here, we provide detailed insight into the mechanism through which GSK3β modulates SIRT2 activity and suggest that the phosphorylation of S327, S331 and S335 may be useful as a target for therapeutic intervention in PD.

#### MATERIALS AND METHODS

#### Materials

An MTT assay kit was purchased from Roche. A site-directed mutagenesis kit was purchased from Stratagene. 6-Hydroxydopamine hydrobromide (6-OHDA), DMSO, SB216763 (S3442, an inhibitor of GSK3β) and AGK2 (A8231, an inhibitor of SIRT2) were obtained from Sigma-Aldrich. Antibodies against pGSK3β (Ser9) and GSK3β were purchased from Cell Signaling (Danvers, MA, USA). Antibodies against SIRT2, ace-tubulin, α-tubulin, HA and Flag were purchased from Sigma-Aldrich. Secondary antibodies conjugated to Alexa 488 or Alexa 594 were purchased from Invitrogen. Hoechst 33258 (94403) was purchased from Sigma-Aldrich. Protein A/G-coated Sepharose beads were obtained from Santa Cruz Biotechnology (Dallas, TX, USA). An antiphosphoserine/threonine/tyrosine antibody was obtained from Abcam (ab15556). Protein kinase CDK5/p25 (cat. 14—516) and GSK3β (cat. 14—306) were purchased from Millipore. Cells were transfected using Lipofectamine 2000 Transfection Reagent (Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA). Other chemicals and reagents were of the highest analytical grade and were purchased from local commercial sources.

#### Cell Culture

The human neuroblastoma cell line SH-SY5Y was obtained from the American Type Culture Collection. The cells were grown in Dulbecco's modified Eagle's medium (DMEM)/Ham's F12 (1:1 mixture; HyClone) supplemented with 10% fetal bovine serum (GIBCO) in a 5% CO2 incubator at 37◦C. Human embryonic kidney cells (HEK293) were grown in DMEM (HyClone) supplemented with 10% FBS.

#### Pharmacological Treatments

6-OHDA was dissolved in phosphate-buffered saline (PBS) and used at a final concentration of 100 µM (Ikeda et al., 2008), which was the dose shown to induce 50% cell death within 24 h after 4 h of exposure. SB216763 and AGK2 were dissolved in DMSO. Before adding 6-OHDA, the SH-SY5Y cells were treated with SB216763 (Acevedo et al., 2014) or AGK2 (Outeiro et al., 2007) at a final concentration of 10 µM for 0.5 h (the DMSO content never exceeded 0.1%).

#### MTT Tests for Cell Viability

Cell viability was measured by a colorimetric MTT [3-(4,5 dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] assay. Briefly, the cells were seeded in a 96-well plate at a density of 5 × 104 cells/well. The cells were then pretreated with SB216763 for 0.5 h. After pretreatment, 6-OHDA was added to the culture medium to reach a final concentration of 100 µM. The control cells were not treated with SB216763 or 6-OHDA. The culture medium was changed after incubation for 4 h. Then, 16 h later, the MTT reagent was added to each well, and the cells were incubated for an additional 4 h. The absorbance of each reaction product was measured with a microplate reader at a wavelength of 595 nm. The results are expressed as a percentage of the MTT absorbance of the control cells, which was set to 100%.

### Western Blot Analysis

Whole-cell lysates were prepared by incubating cells in RIPA buffer supplemented with a protease inhibitor cocktail (Selleck) according to the manufacturer's instructions. Briefly, the cells were harvested by centrifugation at 900 rpm for 5 min and washed in PBS (pH 7.2). The pellets were solubilized in the same volume of lysis buffer, kept on ice, vortexed for 5 min, and centrifuged at 13,000× g for 20 min at 4 ◦C. Equal amounts of total lysate protein were loaded and separated on a 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gel. The proteins were electrophoretically transferred to a PVDF membrane, and the membrane was blocked in 5% skim milk in Tris-buffered saline containing 0.1% Tween-20 (TBST) for 1 h. Then, the membranes were incubated at 4◦C overnight in the presence of a primary antibody against one of the following proteins: HA, Flag, SIRT2, ace-tubulin, α-tubulin (Sigma), GSK3β or pGSK3β (Ser9). Next, the membranes were washed three times with TBST every 5 min and probed with a corresponding horseradish peroxidase (HRP)-conjugated secondary antibody at room temperature for 1 h. Probe detection was conducted using enhanced ECL Advance Western Blotting Detection Reagents (Perkin Elmer, Waltham, MA, USA) and a ChemiDoc XRS+ system (Bio-Rad, Hercules, CA, USA).

#### Immunofluorescence Staining

Cultured cells were fixed with PBS containing 4% paraformaldehyde for 15 min, which was followed by washing and permeabilization in PBS with 0.1% Triton X-100 for 5 min. The samples were blocked in PBS with 5% BSA and incubated with primary antibodies overnight at 4◦C. The following primary antibodies were used: rabbit anti-SIRT2 (1:100) and mouse anti-acetylated α-tubulin (1:50). Secondary antibodies conjugated to Alexa 488 or Alexa 594 (1:500) were applied for 1 h at room temperature. After washing with PBS and staining with DAPI for 5 min, coverslips were mounted with Mowiol. Images of the cells were acquired at room temperature on a confocal microscope (A1+; Nikon) using either a 40× or a 60× lens. Photoshop CS (Adobe) was used to adjust the contrast and brightness.

#### Coimmunoprecipitation

Cells were collected and lysed in buffer containing 50 mM Tris-HCl at pH 7.4, 150 mM NaCl, and 1% NP-40. All buffers were supplemented with protease and phosphatase inhibitors. The extracted proteins were precleared by incubating the lysates with Protein A/G-coated Sepharose beads for 1 h at 4◦C, and then the supernatants (500 µg, 1 µg/µl) were incubated overnight with an HA or Flag antibody at 4◦C. Precipitation of the immune complexes was performed with Protein A/G-coated Sepharose beads for 2 h at 4◦C. After immunoprecipitation, the beads were washed five times with lysis buffer at 4◦C and suspended in 40 µl of 2× loading buffer. All samples were loaded onto a gel, processed by SDS–PAGE and analyzed by Western blot. To detect SIRT2 phosphorylation, we used an antiphosphoserine/threonine/tyrosine antibody.

#### Plasmids and Transfection

GSK3β-HA and SIRT2-flag were obtained from Addgene. PCMV-HA and PCMV-Tag4A are blank plasmids that contain an HA tag and a Flag tag, respectively. A subsequent LR-recombination reaction with PGEX4T3 and phMGFP plasmids resulted in GST-tagged and GFP-tagged expression constructs. Site-directed mutagenesis was performed using a Quick change protocol to generate the S327A, S331A, S335A, S327&331&335A and S327&331&335D mutants. All constructs were verified by DNA sequencing. SIRT2-targeting shRNAs using previously identified sequences (Si et al., 2013) were cloned into a pGPU6/GFP/Neo vector from Genepharma. For transient transfection, cells were transfected using Lipofectamine 2000 Transfection Reagent according to the manufacturer's instructions.

### Determining the Survival Rate of Transfected Cells

SH-SY5Y cells were transfected with plasmids (GFP, GFP-SIRT2, GFP-SIRT2SA or GFP-SIRT2SD) for 24 or 48 h to induce the expression of GFP. Chromatin condensation was detected by nuclear staining with Hoechst 33258. Briefly, the cells were fixed with 4% paraformaldehyde for 15 min, stained with PBS/0.1% TritonX-100/10 µM Hoechst 33258 for 5 min, and then visualized by fluorescence microscopy. Apoptotic cells were stained bright blue because of their chromatin condensation. Images were captured from different fields of each well for different groups of cells at 20× magnification using an Olympus fluorescence microscope. The numbers of total and live transfected cells were counted from multiple fields to obtain the percentage of live transfected cells (Dutta et al., 2018).

#### GST Fusion Protein Purification

The GST-SIRT2 expression plasmid was transformed into competent Escherichia coli BL21 cells. A single colony was inoculated in LB medium at 37◦C in an orbital shaker incubator until the mid-log phase. IPTG (100 µM) was added to the culture and incubated at 20◦C for 8 h. Following the incubation, the culture was centrifuged at 2,500× g for 10 min at 4◦C, and the resulting pellet was resuspended in binding buffer [50 mM Tris-Cl, pH 8.0, 150 mM NaCl and 1× protease inhibitor cocktail (Biotool)]. The cells were lysed by sonication and centrifuged at 12,000× g for 30 min at 4◦C. After centrifugation, the supernatant was collected, incubated with GST agarose beads (Novagen) and maintained under agitation overnight at 4◦C. After overnight binding, the beads were washed with wash buffer (150 mM NaCl, 50 mM Tris-Cl, pH 7.5 and 1% Triton X-100), and the bead-bound proteins were eluted and stored in 30% glycerolcontaining buffer. Other GST fusion proteins were obtained in the same way.

#### In vitro Kinase Assays

The phosphorylation of GST-SIRT2 was performed in a final volume of 25 µl consisting of 5 mM MOPS, pH 7.2, 0.05 mM DTT, 4 mM MgCl2, 80 µm ATP, 10 µCi of [γ-32P]ATP, 0.4 mM EDTA, 1 mM EGTA, and 2.5 mM β-glycerophosphate for 30 min at 30◦C in the presence or absence of CDK5/p25 or GSK3β. The reaction was terminated by the addition of 25 µl of 2× sodium dodecyl sulfate (SDS) loading buffer (187 mm Tris-HCl, pH 6.8, 30% (w/v) glycerol, 6% SDS and 15% β-mercaptoethanol). The samples were then heated at 100◦C for 5 min, centrifuged in a microcentrifuge, and loaded on a gel for SDS-PAGE. Following electrophoresis, the proteins were exposed to a Kodak X-ray film for autoradiography at −80◦C for 2 or 3 days.

(D) Western blot analysis showing SIRT2 phosphorylation in cells. Endogenous SIRT2 was immunoprecipitated from SH-SY5Y cells treated with or without 100 µM 6-OHDA for 4 h and probed with an anti-phosphoserine/threonine/tyrosine antibody to detect phosphorylation levels (∗p < 0.05, ∗∗p < 0.01, two-tailed Student's t-test). The values are expressed as the mean ± SD, n ≥ 3 for each group.

## Phosphopeptide Identification by Mass Spectrometry (MS)

After in vitro phosphorylation, GST-SIRT2 was separated via SDS-PAGE. The gel was cut out after Coomassie blue staining, and the samples were sent to the Beijing Proteome Research Center (BPRC, China).

#### Statistics and Reproducibility

Statistical analysis was performed using Student's two-tailed unpaired t-tests for two-group comparisons and one-way analysis of variance (ANOVA) or two-way ANOVA for multigroup comparisons. P values < 0.05 were considered statistically significant and are indicated with asterisks ( ∗P < 0.05; ∗∗P < 0.01; ∗∗∗P < 0.001; ∗∗∗∗P < 0.0001) in the figure legends. All of the data collected met the normal distribution assumptions of the tests. The data are represented as the mean ± SD. Each experiment was performed at least three times and in duplicate or more.

## RESULTS

### 6-OHDA Elevates the Phosphorylation of SIRT2 in SH-SY5Y Cells

To investigate how toxins regulate SIRT2 function, we first established a cellular model of PD by applying 100 µM 6-OHDA to SH-SY5Y cells at the indicated times and concentrations and then assessed cell viability by the MTT assay. Cell viability decreased in a time-dependent manner following 6-OHDA exposure (**Figure 1A**). Further, we evaluated the expression of SIRT2 in SH-SY5Y cells after treatment with 6-OHDA, and SIRT2 levels showed a slight but not noticeable decrease. Furthermore, the active status of GSK3β was increased (**Figure 1B**). Because SIRT2 can be distributed in the cytoplasm or nucleus under different conditions (Dryden et al., 2003; North et al., 2003; Liu et al., 2017), we wanted to know where SIRT2 is located under 6-OHDA treatment. We next treated SH-SY5Y cells with PBS as a control or 100 µM 6-OHDA for 4 h; later, the location of endogenous SIRT2 was assessed by immunofluorescence. We observed that SIRT2 resided in the cytoplasm for the duration of the experiment (**Figure 1C**). We next purified endogenous SIRT2 from SH-SY5Y cells treated with or without 6-OHDA and detected SIRT2 phosphorylation using an anti-phosphoserine/threonine/tyrosine antibody. We found that phosphorylated SIRT2 levels were significantly increased after treatment with 6-OHDA (**Figure 1D**).

### The Protective Effect of GSK3β Inhibition Occurs Through a Blockade of SIRT2 Phosphorylation

Given that the activity of GSK3β was increased in SH-SY5Y cells treated with 6-OHDA, we blocked the activity of GSK3β in SH-SY5Y cells with SB216763 to determine whether this could protect the cells from 6-OHDA. We pretreated SH-SY5Y cells with different concentrations of SB216763 for 30 min and then applied 100 µM 6-OHDA for 4 h. After that, we measured cell viability and found that SB216763 rescued the SH-SY5Y cells from 6-OHDA (**Figure 2A**). We further analyzed the phosphorylation of endogenous SIRT2 among the control, 6-OHDA and 6-OHDA plus SB216763 groups and found that the phosphorylation of SIRT2 was increased in the 6-OHDA-treated cells, while SB216763 blocked the increase in SIRT2 phosphorylation (**Figure 2B**). These results indicate that GSK3β activation may be responsible for the increased phosphorylation of SIRT2.

SIRT2 is a tubulin deacetylase, and SB216763 increases the acetylation of tubulin (**Figure 2C**), which indicates that the decrease in SIRT2 phosphorylation mediated by SB216763 may reduce SIRT2 deacetylase activity. To further confirm this, we pretreated SH-SY5Y cells with DMSO, SB216763 or AGK2 and monitored the change in acetylated tubulin under 6-OHDA treatment. Acetylated tubulin levels decreased under 6-OHDA treatment, and both SB216763 and AGK2 reversed the decrease in acetylated tubulin levels (**Figure 2D**). To confirm that GSK3β participates in the regulation of SIRT2 deacetylase activity, we overexpressed PCMV-HA or GSK3β-HA in HEK293 cells and treated them with DMSO, SB216763 or AGK2 24 h later. The overexpression of GSK3β resulted in a decrease in acetylated tubulin levels, while SB216763 and AGK2 reversed the decrease in acetylated tubulin levels induced by the overexpression of GSK3β (**Figure 2E**). GSK3β further reduced acetylated tubulin levels relative to SIRT2 overexpression (the acetylated tubulin levels decreased by 23.73% and 40.65% in the absence or presence of GSK3β, respectively; **Figure 2F**). To further confirm that SIRT2 is the downstream target of GSK3β, we used shRNA to knock down endogenous SIRT2 (**Figure 2G**). We found that downregulated SIRT2 could reduce the toxicity of 6-OHDA, while SB216763 could not further improve cell viability after SIRT2 knockdown (**Figure 2H**). These data suggest that SIRT2 is the downstream target of GSK3β under 6-OHDA treatment. Taken together, our data demonstrate that 6-OHDA or the overexpression of GSK3β induces a decrease in acetylated tubulin levels and that inhibiting GSK3β with SB216763 counteracts this effect.

### GSK3β Can Phosphorylate SIRT2 at Three Particular Sites

Given that inhibiting GSK3β blocked an increase in SIRT2 activity, we next tested whether SIRT2 interacts with GSK3β by coimmunoprecipitation (**Figures 3A,B**). Our results indicate that GSK3β and SIRT2 may interact in cells.

To test whether GSK3β directly phosphorylates SIRT2, we carried out an in vitro kinase assay. We purified GST-SIRT2 from E. coli BL21. Because it has been reported that SIRT2 is a phosphorylation substrate for CDK5, we used CDK5/p25 (p25 continues to activate CDK5) as a positive control. We found that GSK3β could phosphorylate SIRT2 (**Figure 3C**). To determine which sites are phosphorylated by GSK3β, we carried out an in vitro kinase assay followed by SDS-PAGE and Coomassie blue staining to detect phosphorylated SIRT2, which was later evaluated by MS (**Supplementary Figure S1**). Finally, we identified three sites that could be phosphorylated by GSK3β (**Figure 3D**). To validate the MS results, we mutated these three sites to alanine residues to mimic dephosphorylation. The mutants were purified from E. coli and subjected to the kinase assay. We found that each mutant decreased the phosphorylation of SIRT2 by GSK3β (**Figure 3E**). To further test whether the interaction between GSK3β and SIRT2 changes under 6-OHDA stress, we performed a coimmunoprecipitation assay. It was shown that the interaction between GSK3β and SIRT2 was increased under 6-OHDA treatment (**Figure 3F**). In conclusion, we identified S327, S331 and S335 as SIRT2 phosphorylation sites targeted by GSK3β.

### SIRT2 Phosphorylation-Resistant Mutants Can Alleviate SIRT2 Toxicity in SH-SY5Y Cells

To further prove that the phosphorylation of SIRT2 by GSK3β does participate in 6-OHDA-induced SH-SY5Y cell death and that overactivated SIRT2 aggravates PD (Outeiro et al., 2007), we constructed two mutants, GFP-SIRT2SA and GFP-SIRT2-SD, to mimic dephosphorylation and phosphorylation, respectively (**Figure 4A**). Given that ace-tubulin is the substrate of SIRT2 (North et al., 2003), we measured the fluorescence intensity of ace-tubulin after the individual overexpression of GFP, GFP-SIRT2, GFP-SIRT2SA or GFP-SIRT2SD, and we found that the mutant that mimicked phosphorylation further decreased the acetylation of α-tubulin, while the mutant that mimicked dephosphorylation did not (**Figures 4B,C**). This result indicates that the phosphorylation-resistant mutant (GFP-SIRT2SA) blocks SIRT2 deacetylation activity. As we showed that GSK3β regulates the phosphorylation of SIRT2 and that the protective effect of blocking GSK3β in the PD cell model occurs through decreased SIRT2 phosphorylation,

FIGURE 2 | Inhibition of GSK3β protected SH-SY5Y cells from 6-OHDA by decreasing SIRT2 phosphorylation. (A) SH-SY5Y cells were pretreated with different concentrations of SB216763 for 30 min and then treated with or without 100 µM 6-OHDA for 4 h. Cell viability was measured by the MTT assay (∗p < 0.05, ∗∗∗∗p < 0.0001, one-way ANOVA followed by Dunnett's post hoc test). (B) Endogenous SIRT2 was immunoprecipitated from whole-cell lysates, and p-SIRT2 was detected by an anti-phosphoserine/threonine/tyrosine antibody (∗p < 0.05, ∗∗p < 0.01, one-way ANOVA followed by Tukey's post hoc test). (C) Western blot analysis showing the acetylation of α-tubulin (ace-tubulin) and total α-tubulin levels in SH-SY5Y cells treated with DMSO or SB216763 at the indicated concentrations (∗p < 0.05, one-way ANOVA followed by Dunnett's post hoc test). (D) SH-SY5Y cells were pretreated with DMSO, 10 µM SB216763 or 10 µM AGK2 for 30 min and then treated with or without 100 µM 6-OHDA for 4 h. Ace-tubulin and α-tubulin were detected by Western blotting (∗∗∗p < 0.001, two-way ANOVA followed by Bonferroni's post hoc test). (E) HEK293 cells were treated with DMSO, 10 µM SB216763 or 10 µM AGK2 for 30 min after transfection with PCMV-HA as a control or GSK3β-HA for 24 h. Ace-tubulin and α-tubulin were detected by Western blotting (∗∗∗p < 0.001, two-way ANOVA followed by Bonferroni's post hoc test). (F) HEK293 cells were cotransfected with GFP (or GFP-SIRT2) and PCMV-HA (or GSK3β-HA) for 24 h. Ace-tubulin and α-tubulin were detected by Western blotting (∗p < 0.05, ∗∗p < 0.01, two-way ANOVA followed by Tukey's post hoc test). (G) The level of the SIRT2 protein was downregulated in SH-SY5Y cells transfected with SIRT2 shRNA (shSIRT2-1 or shSIRT2-2) compared to cells transfected with the control (shNC). SIRT2 and GAPDH were detected by Western blotting. (H) A histogram representing the percentage of viable cells. Cells were transfected with shRNA for 48 h and then treated with 6-OHDA for 4 h. The percentage of viable cells was calculated after 24 h (∗p < 0.05, ∗∗p < 0.01, ns, not statistically significant; one-way ANOVA followed by Tukey's post hoc test). The values are expressed as the mean ± SD, n ≥ 3 for each group.

we next transfected SH-SY5Y cells with the GFP, GFP-SIRT2, GFP-SIRT2SA, or GFP-SIRT2SD plasmids for 24 h or 48 h. Cell viability was measured by live cell counting, and the data showed a notable increase in the viability of the cells overexpressing GFP-SIRT2SA (the GFP-SIRT2 overexpressing cells relative to the GFP-SIRT2SA-overexpressing cells, ∗P < 0.05, ∗∗P < 0.01), whereas there was no increase or decrease in the viability of the cells overexpressing GFP-SIRT2SD (the GFP-SIRT2-overexpressing cells relative to the GFP-SIRT2SD-overexpressing cells, ∗P < 0.05, ∗∗∗P < 0.001; **Figure 4D**). To assess whether AGK2 is able to block phospho-SIRT2 toxicity, we overexpressed the indicated plasmids and found that AGK2 was able to block GFP-SIRT2 toxicity but not GFP-SIRT2SD toxicity (**Figure 4E**).

These data indicate that the phosphorylation of SIRT2 by GSK3β at S327, S331 and S335 accelerates cell death. All of the data suggest that the dephosphorylation of S327, S331, and S335 decreases the activity of SIRT2, which could slow the cell death induced by SIRT2 (GFP-SIRT2) overexpression in SH-SY5Y cells.

#### DISCUSSION

SIRT2 appears to play a detrimental role in neurological disorders such as PD (Outeiro et al., 2007; Chen et al., 2015), Huntington's disease (Luthi-Carter et al., 2010) and ischemic stroke (Krey et al., 2015; Xie et al., 2017; She et al., 2018; Wu et al., 2018), and it is important to understand how SIRT2 is regulated since we know little about it. In the present study, we found that SIRT2 phosphorylation was increased in a cellular model of PD and that this change was mediated by the activation of GSK3β. We further identified three phosphorylation sites in SIRT2 that are targeted by GSK3β and found that a genetic mutant that mimics the dephosphorylation of SIRT2 can alleviate cell death induced by SIRT2 overexpression. Several types of evidence support this finding: (i) using Western blot analysis, we found that the phosphorylation of endogenous SIRT2 in SH-SY5Y cells is increased after treatment with 6-OHDA; (ii) SB216763, an inhibitor of GSK3β, reduced the phosphorylation of endogenous SIRT2; (iii) an in vitro kinase assay showed that SIRT2 is phosphorylated by GSK3β. Phosphoproteomic analysis of SIRT2 revealed that three particular sites are responsible for this phosphorylation; and (iv) genetic mutants that mimic the phosphorylation or phosphorylation resistance of the three phosphorylation sites aggravates or alleviates SIRT2-induced cell death in SH-SY5Y cells, respectively.

In this study, we first determined the subcellular location of SIRT2 under 6-OHDA treatment and found that it remains in the cytoplasm. SIRT2 regularly localizes to the cytoplasm (North et al., 2003), and it has also been reported to localize to the nucleus during the G2/M phase (Dryden et al., 2003; North et al., 2003). This prompted us to search for an upstream regulator of SIRT2 in the cytoplasm. A previous study showed that the acetylation of GSK3β increases in pathological

cardiac hypertrophy and that SIRT2 binds to, deacetylates and activates GSK3β (Sarikhani et al., 2018). SIRT2 may also act as an upstream regulator of GSK3β to modulate the differentiation of dopaminergic neurons (Szegö et al., 2017). Here, we proposed that the increase in SIRT2 phosphorylation mediated by GSK3β affects SIRT2 catalytic activity as well. To obtain the data presented here, we used a mutant that mimics the dephosphorylation of SIRT2 (GFP-SIRT2SA) and found that it does not decrease α-tubulin acetylation, which is the opposite of what is observed with the mutant that mimics the phosphorylation of SIRT2 (GFP-SIRT2SD). Compared to GFP-SIRT2, GFP-SIRT2SA significantly increased the survival of SH-SY5Y cells. This result coincides with that of a previous study (Outeiro et al., 2007). We showed that SIRT2 phosphorylation increases after exposure to 6-OHDA, and we speculated that SIRT2 phosphorylation accelerates cell death. As expected, cell death worsened in the GFP-SIRT2SD group. Thus, the mutant that mimics the dephosphorylation of SIRT2 (GFP-SIRT2SA) can lessen the cell death induced by SIRT2 overexpression by decreasing SIRT2 deacetylase activity. All of these data suggest that there are complex regulatory relationships between SIRT2 and GSK3β.

We found that AGK2 could not inhibit GFP-SIRT2SD toxicity. This suggests that the phosphorylation sites we found here may be important for the ligand docking of AGK2. In previous studies, decreased enzymatic activity was observed in several C-terminal alanine substitution mutations, such as mutations at S327 or S331 and S335 (Nahhas et al., 2007). Interestingly, S331A and S335A individually did not affect SIRT2 enzymatic activity, but double substitutions elicited a 44% reduction in activity (Nahhas et al., 2007). These reports are consistent with our finding that GFP-SIRT2SA reduces SIRT2 deacetylation activity. In contrast, the phosphorylation of S331 or a phospho-mimicking mutant of this site also inhibits the catalytic activity of SIRT2 (Pandithage et al., 2008). This report differs from our finding that GFP-SIRT2SD does not reduce SIRT2 deacetylation activity. This may be due to the simultaneous substitution of three serine residues in SIRT2. A preferred site for ligand binding of AGK2 is the ''C-pocket'' with a hydrogen-bonding pattern that mimics the action of nicotinamide, a known inhibitor of sirtuins (Outeiro et al., 2007). Since S327, S331 and S335 are located outside of the ''C-pocket,'' these serine residues may affect the SIRT2 enzymatic activity by altering the conformation of the protein. In fact, nicotinamide blocks the activity of the phospho-mimicking mutant GST-SIRT-S331D (Pandithage et al., 2008), suggesting that AGK2 also inhibits the activity of GST-SIRT2-S331D. Similarly, we observed that AGK2 does not inhibit GFP-SIRT2SD toxicity, which may be because three simultaneous substitutions alter the conformation of the ''C pocket,'' resulting in the failure of AGK2 docking. Our results indicate that S327, S331 and S335 can be simultaneously phosphorylated by GSK3β in vitro, and we are not sure whether this modification occurs in vivo. More experiments are needed to study this phenomenon.

Interestingly, it has been reported that CDK5 phosphorylates SIRT2 at S331 (Pandithage et al., 2008), S331 and S335 (Zhang et al., 2018), and we found that S327, S331 and S335 are phosphorylated by GSK3β. Furthermore, the phosphorylation of SIRT2 was found to be stronger after the coapplication of CDK5 and GSK3β (data not shown). These results indicate that there may be crosstalk between the GSK3β and CDK5 pathways. The data we present here suggest that phosphorylation-resistant mutants inhibit the catalytic activity of SIRT2. Nevertheless, phosphorylation by CDK5 inhibits the catalytic activity of SIRT2 (Pandithage et al., 2008). S331 is a common phosphorylation site, and the opposite effect may rely on other phosphorylation sites. Thus, targeting CDK5 and GSK3β at the same time may be a potential direction for the development of therapeutic strategies for PD. One possible approach is to block CDK5 and GSK3β simultaneously with one or two inhibitors. Using this strategy, we can obtain inactivated SIRT2 that is unphosphorylated at S327, S331 and S335. This may be the most effective strategy to suppress SIRT2 expression (Nahhas et al., 2007). There are many inhibitors currently in clinical trials that can block CDK5 and GSK3β simultaneously, such as AZD5438 and brain permeable AZD1080. Roscovitine and CHIR-99021, which are inhibitors of CDK5 and GSK3β, respectively, can also be used together. However, it is worth noting that the activities of CDK5 and GSK3β are related and that crosstalk depends on age (Engmann and Giese, 2009). In young mice, an increase in p25-induced CDK5 activity inhibits GSK3β activity by enhancing inhibitory phosphorylation at Ser-9 of GSK3β (Plattner et al., 2006). However, in older mice, GSK3β activity is enhanced in p25 transgenic mice (Plattner et al., 2006; Wen et al., 2008). Thus, the reaction of young and aged mice to the simultaneous targeting of CDK5 and GSK3β may be different.

As aging is considered a major risk factor for the development of PD, the pathways involved in aging may

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Acevedo, K. M., Opazo, C. M., Norrish, D., Challis, L. M., Li, Q. X., White, A. R., et al. (2014). Phosphorylation of amyloid precursor protein at threonine 668 is essential for its copper-responsive trafficking in SH-SY5Y provide targets for therapeutic interventions for PD. SIRT2 is a member of the sirtuin family of proteins, which participate in a variety of cellular functions and play a role in aging (Maxwell et al., 2011). Outeiro et al. (2007) found that the inhibition of SIRT2 rescues α-synuclein toxicity and modifies inclusion morphology in a cellular model of PD (Dillin and Kelly, 2007; Garske et al., 2007). Later, de Oliveira et al. (2017) showed that α-synuclein is deacetylated by SIRT2 and that α-synuclein acetylation is a key regulatory mechanism governing α-synuclein aggregation and toxicity. Here, we further characterized SIRT2, showing that its activity is activated by its phosphorylation by GSK3β in a cellular model of PD. Based on the above conclusions, we elucidated a complete GSK3β-SIRT2-α-synuclein signaling cascade that participates in the development of PD. Because of the importance of α-synuclein in PD, we believe this cascade may play a vital role in PD. We may find treatments that inhibit the progression of PD by targeting this cascade.

Taken together, the data presented here highlight a previously unexplored cellular pathway that might underlie the impairment of dopaminergic neurons in PD, further underscoring the potential of the GSK3β-SIRT2-α-synuclein signaling cascade as a viable target pathway for neuroprotective therapies.

#### AUTHOR CONTRIBUTIONS

XW designed the experiments. SL and ZZ performed the enzyme activity assays and the biochemical analysis of the SH-SY5Y cells. LZ, SM and SLi analyzed the data. SL and ZZ wrote the manuscript with help from XW. All authors read and approved the final manuscript.

#### FUNDING

This work was supported by the Natural Science Foundation of Guangdong (2016A030311009), the National Natural Science Foundation of China (31371184), and the Program for Changjiang Scholars and the Innovative Research Team in University (IRT\_16R37).

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fncel.2019.001 48/full#supplementary-material

FIGURE S1 | Phosphopeptide identification by mass spectrometry (MS). (A) GST-SIRT2 was purified from BL21 E. coli, and an in vitro kinase assay was performed with or without GSK3β. The samples were separated via SDS-PAGE, and then the gel was visualized with Coomassie blue staining. The band in the red rectangle was subjected to MS. (B) The serine residues in red are the predicted phosphorylated sites.

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Chen, X., Wales, P., Quinti, L., Zuo, F., Moniot, S., Herisson, F., et al. (2015). The sirtuin-2 inhibitor AK7 is neuroprotective in models of Parkinson's disease but not amyotrophic lateral sclerosis and cerebral ischemia. PLoS One 10:e0116919. doi: 10.1371/journal.pone. 0116919

**Conflict of Interest Statement**: The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Liu, Zhou, Zhang, Meng, Li and Wang. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Lipopolysaccharide-Induced Autophagy Mediates Retinal Pigment Epithelium Cells Survival. Modulation by the Phospholipase D Pathway

Vicente Bermúdez 1,2 , Paula Estefanía Tenconi 1,2 , Norma María Giusto1,2 and Melina Valeria Mateos 1,2 \*

1 Instituto de Investigaciones Bioquímicas de Bahía Blanca (INIBIBB), Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Bahía Blanca, Argentina, 2Departamento de Biología, Bioquímica y Farmacia (DBByF), Universidad Nacional del Sur (UNS), Bahía Blanca, Argentina

Inflammation and oxidative stress are common factors involved in the pathogenesis of retinal diseases, such as aged-related macular degeneration (AMD) and diabetic retinopathy (DR). Autophagy is a catabolic process essential to cell survival in response to stress. This process is highly active in retinal pigment epithelium (RPE) cells. Our previous findings demonstrated that lipopolysaccharide (LPS) induces an inflammatory response of RPE cells that implies classical phospholipases D (PLD1 and 2) activation, cyclooxygenase-2 (COX-2) expression, prostaglandin E2 (PGE2) production and reduced cell viability. In this work, we studied the autophagic process and its modulation by the PLD pathway in D407 and ARPE-19 RPE cells exposed to LPS. LPS (10 µg/ml or 25 µg/ml) exposure for 24 h increased light chain 3B-II (LC3B-II) content (an autophagy marker) and LC3B-positive punctate structures in both RPE cell lines studied. Next, the drug bafilomycin A1 (BAF, 50 nM) was used to block the autophagic flux. In cells pre-incubated with BAF, LC3B-II and sequestosome 1 (SQSTM1/p62) levels and autophagosome-like structures were increased by LPS, demonstrating that the inflammatory injury increases the autophagic process in RPE cells. To study the role of the PLD pathway, cells were pre-incubated for 1 h with selective PLD1 (VU0359595) or PLD2 (VU0285655-1) inhibitors prior to LPS addition. Under control condition, LC3B-positive punctate structures were increased in cells pre-incubated with PLD2 inhibitor while with PLD1 inhibitor were increased in cells exposed to LPS. MTT reduction assays showed that early autophagy inhibitors, 3-methyladenin (3-MA) or LY294002, enhanced the loss in cell viability induced by LPS exposure for 48 h. On the contrary, the inhibition of PLD1 and PLD2 prevented the loss in cell viability induced by LPS. In conclusion, our results show that even though LPS treatment promotes

Bermúdez V, Tenconi PE, Giusto NM and Mateos MV (2019) Lipopolysaccharide-Induced Autophagy Mediates Retinal Pigment Epithelium Cells Survival. Modulation by the Phospholipase D Pathway. Front. Cell. Neurosci. 13:154. doi: 10.3389/fncel.2019.00154

<EMAIL_ADDRESS><EMAIL_ADDRESS>
Federal University of Rio de Janeiro,

Maria Del Carmen Fernandez, Universidad de Buenos Aires,

Received: 02 January 2019 Accepted: 08 April 2019 Published: 24 April 2019

#### Citation:

Edited by: Mario Eduardo Guido, Center for Research in Biological Chemistry Córdoba (CIQUIBIC),

> Argentina Reviewed by: Rafael Linden,

> > Brazil

Argentina

\*Correspondence: Melina Valeria Mateos

**Abbreviations:** 3-MA, 3-methyladenine; BAF, bafilomycin A1; BECN1, beclin1; COX-2, cyclooxygenase-2; DAG, diacylglycerol; ERK, extracellular signal-regulated kinase; HRP, horseradish peroxidase; LC3, microtubule-associated protein 1 light chain 3; LPS, lipopolysaccharide; LPPs, lipid phosphate phosphatases; mTOR, mammalian target of rapamycin; NFκB, nuclear factor kappa B; RAP, rapamycin; SQSTM1/p62, sequestosome 1; PA, phosphatidic acid; PC, phosphatidylcholine; PGs, prostaglandins; PKC, protein kinase C; PKD, protein kinases D; PLD, phospholipase D; POS, photoreceptor outer segments; PR, photoreceptor; PVDF, polyvinylidene fluoride; RasGRP, Ras guanine-releasing protein; RPE, retinal pigment epithelium; WB, western blot.

an inflammatory response in RPE cells, it also triggers the activation of the autophagic process which in turn may serve as a protective mechanism for the cells. In addition, we demonstrate that the PLD pathway modulates the autophagic process in RPE cells. Our findings contribute to the knowledge of the molecular basis of retinal inflammatory and degenerative diseases and open new avenues for potential therapeutic exploration.

Keywords: retinal pigment epithelium, lipopolysaccharide, autophagy, inflammation, phospholipase D

#### INTRODUCTION

The retinal pigment epithelium (RPE) is a monolayer of pigmented epithelial cells located between the retina and the vascular choroid, constituting the outer blood-retinal barrier (BRB). These epithelial cells play various critical roles for the correct function of the neural retina and photoreceptor (PR) survival, such as the secretion of several growth factors and cytokines and the transport of nutrients and water to the retina. Also, they protect against photooxidation and mediate the re-isomerization of all-trans-retinal and the renewal of photoreceptor outer segments (POS) by phagocytosis (Strauss, 2005; Strauß, 2016; Carr et al., 2009).  Likewise, RPE mediates in the immune response of the retina, by secreting pro- and anti-inflammatory cytokines, chemoattractants proteins, adhesion molecules and complement factors (Strauss, 2005; Strauß, 2016; Simó et al., 2010; Viringipurampeer et al., 2013).