Expression of leukemia inhibitory factor in Müller glia cells is regulated by a redox-dependent mRNA stability mechanism
© Agca et al. 2015
Received: 7 January 2015
Accepted: 8 April 2015
Published: 25 April 2015
Photoreceptor degeneration is a main hallmark of many blinding diseases making protection of photoreceptors crucial to prevent vision loss. Thus, regulation of endogenous neuroprotective factors may be key for cell survival and attenuation of disease progression. Important neuroprotective factors in the retina include H2O2 generated by injured photoreceptors, and leukemia inhibitory factor (LIF) expressed in Müller glia cells in response to photoreceptor damage.
We present evidence that H2O2 connects to the LIF response by inducing stabilization of Lif transcripts in Müller cells. This process was independent of active gene transcription and p38 MAPK, but relied on AU-rich elements (AREs), which we identified within the highly conserved Lif 3′UTR. Affinity purification combined with quantitative mass spectrometry identified several proteins that bound to these AREs. Among those, interleukin enhancer binding factor 3 (ILF3) was confirmed to participate in the redox-dependent Lif mRNA stabilization. Additionally we show that KH-type splicing regulatory protein (KHSRP) was crucial for maintaining basal Lif expression levels in non-stressed Müller cells.
Our results suggest that H2O2-induced redox signaling increases Lif transcript levels through ILF3 mediated mRNA stabilization. Generation of H2O2 by injured photoreceptors may thus enhance stability of Lif mRNA and therefore augment neuroprotective LIF signaling during degenerative conditions in vivo.
Lack of detailed knowledge about molecular disease mechanisms poses a primary challenge for the development of new therapeutic strategies for blinding diseases. One approach to prevent blindness is to stimulate endogenous neuroprotective pathways to support survival of stressed or injured retinal cells. Whereas overexpression of neurotrophic factors delays or inhibits retinal degeneration [1-7], inhibition of neuroprotective signaling or absence of neuroprotective factors, such as leukemia inhibitory factor (LIF), brain derived neurotrophic factor (BDNF) and fibroblast growth factor 2 (FGF2) accelerates retinal degeneration in disease models, or in the aging retina [2,8,9]. Although increased levels of neurotrophic factors are beneficial for injured retinal cells, a balanced expression is required in the normal retina since exaggerated doses may have detrimental side effects [3,10,11].
One of the neuroprotective factors that is tightly regulated in the neuronal retina is LIF. Lif is expressed in a small and dispersed subpopulation of Müller glial cells in response to photoreceptor injury  and signals through the LIFR/gp130 receptor complex activating the Janus kinase (JAK)/signal transducer and activator of transcription 3 (STAT3) signaling pathway [2,3,12,13]. Activation of this pathway leads to increased expression of endothelin-2 (Edn2), Fgf2, Stat3, Jak3, suppressor of cytokine signaling 3 (Socs3) and glial fibrillary acidic protein, (Gfap) [2,12]. Elimination of LIF impairs expression of these factors in the retina and results in a more severe disease progression [2,12]. Thus, LIF induces complicated intercellular signaling events between degenerating photoreceptors and Müller cells that are crucial for photoreceptor survival [2,12-14].
Upregulation of LIF signaling has been observed in induced and inherited photoreceptor degeneration models [2,13,15,16], as well as in models of ganglion cell death [17-19]. Therefore, induction of Lif expression may be a common mechanism in the injured retina to support neuronal survival and may be one of the main tasks of Müller cells in their attempt to protect retinal cells against degeneration. Despite its important role in neuronal survival and its unique expression profile in the injured retina, the molecular mechanisms that regulate Lif expression in Müller cells are only poorly understood. Recently, we showed that activation of Lif gene transcription in the injured retina involves p38 MAPK signaling , but additional regulatory mechanisms are likely to exist.
Previous reports have shown that injured photoreceptors generate H2O2 through nicotinamide adenine dinucleotide phosphate-oxidase (NOX) enzyme complexes [21-23]. In the presence of NOX inhibitors, generation of H2O2 is impaired and photoreceptor apoptosis is increased in the presence of toxic stress [21-23]. Moreover, increased levels of reactive oxygen species (ROS) upregulate extracellular signal regulated kinase (ERK) and v-akt murine thymoma viral oncogene homolog kinase (AKT) dependent pathways and inhibit the activity of protein phosphatase 2 (PP2A), all of which critically affect photoreceptor survival [24,25]. This seems controversial since H2O2 and other ROS are well known to have detrimental effects on cell function and viability, and many reports show that oxidative stress contributes to retinal degenerative diseases [26-29]. However, it is now clear that subtoxic levels of H2O2 have important roles in signal transduction and are involved in many biological pathways [30,31]. Low levels of H2O2 can reversibly oxidize selective amino acids, such as cysteine, histidine, methionine and selenocysteine, and thus modulate molecular pathways associated with such modified proteins [32-38]. Subtoxic doses of H2O2 were also shown to participate in neuroprotection by ischemic preconditioning  and to induce axonal regeneration in zebrafish , supporting the concept that generation of H2O2 has neuroprotective consequences during stress conditions. Therefore, an intriguing hypothesis suggests that H2O2 generated by NOX enzymes or released from mitochondria in stressed cells may act as a physiological messenger to regulate expression of neuroprotective factors in Müller cells. This hypothesis is supported by the previously reported regulation of Lif expression by p38 MAPK , since p38 MAPK signaling can be activated by H2O2 and may interfere with mRNA stability of target genes. This level of gene regulation involves several RNA binding proteins including tristetraprolin (TTP), which is known to be regulated by p38 MAPK itself [41-43].
Here, we show that H2O2 enhanced mRNA stability of Lif during stress in a Müller cell line and in primary mouse Müller cells. Highly conserved AU-rich elements (AREs) in the Lif 3′UTR were important for this regulation and provided target sequences for several RNA binding proteins. Of those, interleukin enhancer binding factor 3 (ILF3) was identified to be critically involved in the regulation of the H2O2-dependent increase of Lif mRNA stability, and KH-type splicing regulatory protein (KHSRP) was identified to be a general regulator of Lif mRNA levels independent of redox signaling. Our results highlight the complex regulation of Lif expression, and provide a mechanism for the puzzling connection between redox signaling and expression of survival factors such as LIF in Müller glia cells.
H2O2 stabilizes Lif mRNA in Müller glia cells
Signaling between degenerating photoreceptors and Müller glia cells induces expression of several neuroprotective factors for photoreceptor survival [2,13-15]. Recent evidence suggests that redox mechanisms may be involved in this intercellular communication, and it was proposed that H2O2, which is produced by stressed photoreceptors, might be a molecule responsible for the induction of retinal survival pathways [22-24].
To test whether a transcriptional or posttranscriptional mechanism was responsible for the H2O2-induced increase of Lif mRNA levels, we used actinomycin D (ActD) to block RNA polymerase II dependent transcription. Like SD, ActD treatment caused a rapid decrease of Lif mRNA levels, whereas other transcripts (except for Ttp) were not strongly affected. Importantly, H2O2 increased Lif transcripts by 2.2 fold (one hour, P <0.05) and 3.2 fold (two hours, P <0.05) also in the presence of ActD (Figure 1B) suggesting a posttranscriptional mechanism for the control of Lif mRNA levels. Since H2O2 is a reactive molecule that may potentially reduce the activity of ActD, we confirmed that ActD was fully active in the presence of H2O2 and completely blocked tumor necrosis factor-alpha (TNF)-induced Lif transcription (Additional file 1: Figure S1). H2O2 also significantly increased Ttp mRNA levels in the presence of ActD (Figure 1B) showing that Ttp mRNA levels can also be regulated on a posttranscriptional level in rMC-1 cells.
Inhibition of p38 MAPK signaling and addition of H2O2 also affected mRNA levels of cyclooxygenase 2 (Cox2) and Ttp comparably to Lif (Figure 3A). Gfap levels, however, were not significantly altered by the treatments. Interestingly, mRNAs of Cox2 and Ttp, but not of Gfap, contain several AREs that regulate their stability [42,54].
Cis-regulation of Lif mRNA stability
Vector list of cloned regions used in constructs
Lif 3′UTR position (1 to 3180)
Length of 3′UTR (bp)
Whole Lif 3′UTR
1 to 3180
AU-rich R I
1 to 570
AU-rich R II
2556 to 3180
2999 to 3180
2805 to 2998
2541 to 2812
2079 to 2245
1062 to 1231
398 to 570
62 to 229
2999 to 3180
2999 to 3180
2999 to 3180
2999 to 3180
1062 to 1231
1062 to 1231
1062 to 1231
1062 to 1231
R27 2 ARE
2999 to 3180
R27 4 ARE
2999 to 3180
R27 9 ARE
2999 to 3180
Fusion of the whole Lif 3′UTR (3,180 bp) to the reporter gene reduced expression of Luc2CP by 89% compared to a construct with a minimal 3′UTR (ΔUTR) (Figure 4B). When the full length Lif 3′UTR was compared to a similar-sized human genomic sequence without conserved, genic or repetitive sequences this reduction was 55% (Figure 4B), excluding that Lif mRNA destabilization was mediated non-specifically by its long 3′UTR sequence. This suggested that the Lif 3′UTR contained cis regulatory elements leading to reduced levels of the reporter protein. Test of the individual AU-rich regions I and II (570 bp and 625 bp, respectively; Table 1) showed that region II strongly reduced luciferase levels by 43%, whereas region I had no effect when compared to a similar sized region from the human ACTB 3′UTR (Figure 4B). This suggested that region II may contain elements for the cis-regulation of Lif mRNA stability. To identify the elements responsible for the effect, we tested shorter sequences within both regions. In addition, short sequence elements that contained a putative binding site for miR-29 in connection with two AREs (region 36) or a putative binding site for miR-17 (region 38) were also tested (Figure 4C, Table 1). Whereas several of these sequence elements decreased luciferase levels only mildly, regions 36 and 27 had strong and significant effects (Figure 4C). These data identified two small regions important for the regulation of Lif mRNA stability and suggested that region 27 may be responsible for the destabilizing effect observed with the larger region II (Figure 4B). To confirm that regions 27 and 36 regulate Lif mRNA levels through RNA stability, we transfected rMC-1 cells with the respective reporter constructs and followed Luc2PC luminescence in the presence of ActD. Cells expressing the reporter gene fused to regions 27 or 36 lost Luc2PC luminescence significantly faster than cells expressing the reporter containing region 32 (no ARE) (Figure 4D). Thus, regions 27 and 36 influenced RNA levels, indeed, through regulation of RNA stability.
Overall, these experiments identified ARE elements within regions 27 and 36 that are involved in the regulation of Lif mRNA stability in Müller cells.
ILF3 influences redox-dependent Lif mRNA stability in Müller glia
List of RNA binding proteins associated with region 27 of the Lif 3′UTR
27 wt versus 27-No Significance
27 wt versus 27-No Rel. levels
ELAV-like protein 1
Translationally-controlled tumor protein
Fatty acid-binding protein, epidermal
Interleukin enhancer-binding factor 3
Citrate synthase, mitochondrial
Prostaglandin E synthase 3
Putative tropomyosin alpha-3 chain-like protein
Interleukin enhancer-binding factor 2
Small ubiquitin-related modifier 2-3-4
Isocitrate dehydrogenase [NADP], mitochondrial
Spermatid perinuclear RNA-binding protein
Cellular nucleic acid-binding protein
Phosphatidylethanolamine-binding protein 1
Protein mago nashi homolog-B
Rab GDP dissociation inhibitor beta
Aspartate aminotransferase, mitochondrial
Acidic leucine-rich nuclear phosphoprotein 32 family member E
Chloride intracellular channel protein 1
GTP-binding protein SAR1a
Thioredoxin domain-containing protein 5
RNA-binding motif, single-stranded-interacting protein 1
Prefoldin subunit 5
Protein disulfide-isomerase A4
Ubiquitin-conjugating enzyme E2 N
Ras-related protein Rap-1A
Fructose-bisphosphate aldolase C
Endoplasmic reticulum resident protein 29
Heterogeneous nuclear ribonucleoproteins A2/B1
Protein disulfide-isomerase A3
Actin-related protein 2/3 complex subunit 5
L-lactate dehydrogenase A chain
Astrocytic phosphoprotein PEA-15
Peptidyl-prolyl cis-trans isomerase A
Macrophage migration inhibitory factor
Heterogeneous nuclear ribonucleoprotein D0
Heterogeneous nuclear ribonucleoprotein A/B
Heterogeneous nuclear ribonucleoprotein D-like
List of RNA binding proteins associated with region 36 of the Lif 3′UTR
36 wt versus 36-No Significance
36 wt versus 36-No Rel. levels
60S ribosomal protein L36
Endoplasmic reticulum resident protein 29
Peptidyl-prolyl cis-trans isomerase FKBP1A
Citrate synthase, mitochondrial
60S ribosomal protein L18a
60S ribosomal protein L35
Leucine-rich repeat-containing protein 59
Serine/arginine-rich splicing factor 2
Protein transport protein Sec61 subunit gamma
Far upstream element-binding protein 1
Tropomyosin alpha-1 chain
ADP-ribosylation factor-like protein 1
Synaptic vesicle membrane protein VAT-1 homolog
60S ribosomal protein L19
Ras-related protein Rab-2A
GrpE protein homolog 1, mitochondrial
Thioredoxin-like protein 1
Threonine--tRNA ligase, cytoplasmic
Fructose-bisphosphate aldolase C
Hydroxymethylglutaryl-CoA synthase, cytoplasmic
Actin, cytoplasmic 2; Actin, cytoplasmic 2, N-terminally processed
DNA-binding protein A
Acidic leucine-rich nuclear phosphoprotein 32 family member E
60S ribosomal protein L3
Translationally-controlled tumor protein
Eukaryotic translation initiation factor 1
60S ribosomal protein L34
Nuclear migration protein nudC
Squamous cell carcinoma antigen recognized by T-cells 3
High mobility group protein B1; Putative high mobility group protein B1-like 1
60S ribosomal protein L13a; Putative 60S ribosomal protein L13a-like
AP-1 complex subunit gamma-1
Transcriptional activator protein Pur-alpha
60S ribosomal protein L10
Phosphoglycerate kinase 1
Ubiquitin-like modifier-activating enzyme 1
Ras-related protein Rab-1B
Ras-related protein Rab-14
Alanine--tRNA ligase, cytoplasmic
PRA1 family protein 3
Heterogeneous nuclear ribonucleoprotein H2
Phosphatidylethanolamine-binding protein 1; Hippocampal cholinergic neurostimulating peptide
Ubiquitin-conjugating enzyme E2 L3
Structural maintenance of chromosomes protein 3
ELAV Like RNA Binding Protein 1 (ELAVL1) and ILF3, two well-known ARE-binding proteins [62-65], were identified as primary candidates for targeting AREs in region 27 of the Lif 3′UTR (Table 2). Interestingly, ILF3 together with ELAVL1 was previously shown to regulate gene expression of mitogen-activated protein kinase phosphatase 1 (Mkp1) through an H2O2-dependent mechanism . ILF3 was additionally shown to interact with several RNA binding proteins including heterogeneous nuclear ribonucleoprotein (HNRNP) D, HNRNPA2/B1 and HNRNPA/B . All of these HNRNPs were identified by our MS analysis and shown to preferentially bind the 27 wild type over the respective ARE-null sequence (Table 2); but none reached statistical significance (P >0.05). Since HNRNPD not only is an interacting partner of ILF3 but also a known ARE binding protein, which potentially directly influences stability of target mRNAs [67-69], we included it in our further analyses.
This argued that redox regulated Lif mRNA stability may involve ILF3 but not ELAVL1 or HNRNPD.
KHSRP is an important general regulator of Lif expression
Regulation of Lif expression in Müller cells
Our results show that Lif mRNA levels are regulated in stressed Müller cells by a redox controlled stability mechanism that involves AREs in the Lif 3′UTR. This mechanism may thus constitute a significant part of the neuroprotective activity of Müller glia cells that critically depends on signaling between injured photoreceptors and Müller cells [2,13-15]. Several reports argue that H2O2 may act as a signaling molecule in various biological systems [31,32]. Our results support this notion and suggest that the reported neuroprotective effect of H2O2 in the retina [22-24] may be through its signaling to Müller cells leading to increased expression of Lif. H2O2 may be generated by NADPH oxidases or be released from outer segments in case of photoreceptor injury . Generation or release may occur as long as stress conditions exist and, thus, H2O2 may signal for a prolonged period of time to ensure a lasting increase of Lif mRNA levels in the damaged retina as observed in both the VPP and rd10 models of retinal degeneration [2,16].
In the mammalian retina, basal expression of Lif is barely detectable but expression increases after photoreceptor or ganglion cell injury [2,17,19,75]. However, in both primary Müller cells and rMC-1 cells basal Lif expression was relatively high, a phenomenon which may be the result of removing Müller cells from their tissue environment disrupting cell-cell interactions and/or of in vitro culture conditions. Still, Lif expression responded to TNF treatment in rMC-1 cells in vitro leading to transiently increased levels similar to the in vivo situation . Importantly, H2O2 also enhanced Lif mRNA stability after such a transient upregulation of Lif transcription (Figure 2) highlighting the importance of H2O2 signaling for the prolonged maintenance of increased Lif expression during stress conditions. Under normal non-stress conditions, however, H2O2 did not affect Lif levels (Additional file 1: Figure S1), in contrast to mitogen-activated protein kinase phosphatase 1 (Mkp1) and placenta growth factor (Plgf) that were induced under such conditions [46,64]. Because of its reported side effects in the retina [3,11,12,76], Lif expression might be needed to be tightly regulated by a negative feedback system to avoid hazardous high LIF levels under normal conditions. Thus, H2O2 may be required for Lif to overcome this negative feedback and to sustain Lif RNA stability ultimately leading to an increased cell survival during degenerative conditions in the retina.
An additional level of control may be the restriction of Lif upregulation to a small subpopulation of Müller cells in the injured retina . This may additionally ensure that overall levels of LIF may not exceed a certain threshold, thus safeguarding the retina. Although the molecular mechanism of this regulation is not known, an intriguing possibility is the existence of programmed Müller cells that have the unique capability to respond to H2O2 as a physiological messenger. Recently, aquaporins 3 and 8 have been identified as channels for H2O2, a molecule once believed to diffuse freely through cell membranes [47,53,77-80]. Although all known aquaporins have been identified in ocular tissues  and aquaporin-4 has been shown to alter its localization in Müller glia cells upon photoreceptor injury , it will be important to study the aquaporin expression profiles and localization with respect to H2O2 signaling and the ensuing Müller cell response.
Even though H2O2 clearly functioned through the regulation of Lif mRNA stability without affecting Lif gene transcription, H2O2 may nevertheless act on other levels as well. It has been shown that H2O2 activates expression of nuclear factor erythroid 2-related factor (NFE2L2) target genes such as sulfiredoxin and heme-oxygenase 1 . NFE2L2 is a redox regulated transcription factor prominently expressed in Müller cells and astrocytes in the retina [8,83] and binds to specific sequence elements in the 5′UTR of target genes. Interestingly, such elements were not only identified in the Lif 5′UTR, but they also had the highest cross-species conservation among other transcription factor binding sites . This raises the possibility that a connection might exist between H2O2 levels, NFE2L2 activation and initial Lif expression. However, we have not addressed the initiation of Lif transcription in this study as Lif expression was already high in cultured Müller cells. Therefore, in vivo studies are required to study such a potential interaction and to better understand redox regulation of Lif expression during retinal disease conditions.
Cis-acting elements for Lif regulation
Various studies have shown that regulation of mRNA stability and, hence, gene expression is closely linked to regulatory sequence elements within the 3′UTR of the target gene. Among those, AREs regulate stability of the associated gene transcript by directing the binding of regulatory proteins. Hao et al. have shown that the number of AREs is closely associated with the timing of events during an inflammatory response. Early response genes have the highest number of AREs, which may support fast turnover of transcripts after the initial activation to strictly regulate gene expression . Lif mRNA is also associated with a high number of AREs in its 3′UTR (Additional file 3: Figure S3A) and has been shown by us and others to be expressed early and transiently in response to injury [17,21,85]. AREs that were associated with the regulation of Lif mRNA stability resided mainly in highly conserved regions in mammals including humans (Figure 4A). This suggests that mechanisms for the regulation of Lif expression may be conserved among mammals.
Since H2O2 increased mRNA levels not only of Lif but also of other ARE containing transcripts such as Ttp and Cox2 (Figure 3A), our data support the concept that H2O2 signaling may control a general pathway for the regulation of ARE-mediated mRNA stability during oxidative stress. Additionally, H2O2 signaling may also be linked to inflammation since (1) TTP can regulate expression of critical inflammatory response genes such as Tnf [41,43], (2) COX2 is an important mediator of the inflammatory response , and (3) Lif signaling has both pro- and anti-inflammatory properties [87-90]. Since inflammatory events have been implicated in retinal pathologies including age related macular degeneration (AMD) , H2O2 signaling may not only be neuroprotective via regulation of Lif expression but may affect the outcome of retinal degenerative diseases on several levels.
Trans-acting factors for Lif regulation
MS analysis has identified several RNA binding proteins that targeted Lif ARE sequences. ELAVL1 and ILF3 were of significant interest among the proteins that bound to AREs of region 27, as both proteins were previously shown to be involved in H2O2 dependent mRNA stabilization and/or translation . Here, ILF3 but not ELAVL1 was involved in redox dependent regulation of Lif mRNA stability in Müller cells (Figure 6). This suggests that ELAVL1 either may not be involved in the regulation of Lif mRNA stability or may influence Lif expression by other means such as through regulation of RNA splicing [92,93] and/or translation .
Interestingly, neither Lif nor Mkp1 mRNAs were identified as ILF3 target RNAs by a high-throughput ribonucleoprotein immunoprecipitation assay  even though both Lif (this study) and Mkp1 (Kuwano et al. ) were shown to be regulated by ILF3 during oxidative stress. Since non-treated HeLa cell lysates were used for those experiments , this may indicate that ILF3 may have different binding affinities for target sequences under normal and stress conditions. Here, however, ILF3 was identified as a binding protein for AREs in the Lif 3′UTR using extracts from untreated Müller cells (Table 2) and shown to function under redox conditions. Although experimental approaches and binding conditions differed from the study of Kuwano , this indicates that the binding affinity of ILF3 to target AREs may be influenced by cell-type and/or species specificities.
In contrast to ILF3, the contribution of KHSRP to the regulation of Lif mRNA expression in Müller cells may not depend on redox signaling (Figure 7). Similar to the inhibition of p38 MAPK, siRNA-mediated knockdown of Khsrp severely reduced Lif mRNA levels suggesting that KHSRP is required to stabilize Lif transcripts under normal conditions. This result was unexpected, since KHSRP has been shown to negatively regulate stability of its target mRNAs [71,96]. Although KHSRP was detected in our MS analysis, it did not show preferential binding to either wild type or mutant sequences (dataset PXD001463 on the ProteomeXchange Consortium platform ). Thus, it is possible that the reduction of Lif mRNA levels in the absence of KHSRP may be independent of AREs in regions 27 and/or 36 of the 3′UTR. Since KHSRP has been shown to promote maturation of miRNAs , KHRSP may affect Lif mRNA levels indirectly, potentially by modulating non-coding RNAs involved in Lif regulation. Although miRNAs are generally believed to destabilize target mRNAs, evidence has been presented that individual miRNAs may also increase the stability of targets . Thus, absence of KHSRP may prevent maturation of specific non-coding RNAs required for Lif mRNA stabilization under normal conditions. Alternatively, KHSRP may act through an interaction with the p38 MAPK pathway  since both inactivation of KHSRP and inhibition of p38 MAPK had similar consequences for Lif mRNA levels under normal conditions.
Our results highlight important aspects of Lif gene expression, which may impact on retinal physiology and pathophysiology. Specifically, regulatory proteins identified here may provide attractive targets for the modulation of LIF synthesis in retinal degenerative diseases since a moderate elevation of endogenous Lif expression may be neuroprotective and support photoreceptor survival (CA, unpublished data). Our data may also impact on stem cell biology since Lif is a pleiotropic factor that contributes to stem cell renewal in vivo and maintenance of pluripotency of mouse stem cells in culture [98,99]. Interestingly, Müller cells act as stem cells to regenerate retinal neurons in fish  and attempt (but fail) to do likewise in mouse [101-103]. Thus, a proper adjustment of Lif signaling in vivo may influence the capability of mammalian Müller cells to act as progenitors in retinal disease. Clearly, in vivo experiments are warranted to test whether modulating the components of LIF regulation may affect neuroprotection and/or impact on the stem cell potential of Müller glia cells in the mouse retina and may thus improve disease outcome and vision.
Neuroprotection through the generation of ROS by injured neurons has long been a controversial issue in neurodegenerative diseases. Here, we show that H2O2 can act as a messenger to regulate expression of the neuroprotective gene Lif in stressed Müller glia. Redox dependent increase in expression is achieved by the modulation of Lif mRNA stability through an Ilf3 dependent pathway and conserved AREs in the 3′UTR. Therefore, injury-induced production of ROS by retinal neurons leads to stabilization of Lif mRNA that may result in a more sustained expression when LIF signaling is necessary to preserve neurons.
Cell culture, serum deprivation, H2O2, ActD, TNF and p38 MAPK inhibitor treatment
The rat Müller glia cell line rMC-1  was obtained from Dr. Sarthy (Northwestern University, Chicago, USA, IL). Cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM; Life Technologies, Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS; Life Technologies), 100 U/ml penicillin and 100 μg/ml streptomycin (Life Technologies), and grown in a humidified 5% CO2 incubator as described . A total of 300,000 cells in 2 ml growth media were seeded on a six-well plate and cultured overnight. For serum deprivation, rMC-1 cells were washed once with warm PBS and growth media without serum was added. Then, 30% H2O2 (Sigma Aldrich, St. Louis, MO, USA) was diluted in DMEM with or without FBS to a final concentration of 50 μM and added to PBS-washed cells. ActD (Sigma Aldrich) was diluted in DMEM with or without FBS/H2O2 to a final concentration of 10 μg/ml. Rat recombinant TNF (R&D systems, Minneapolis, MN, USA) and the p38 MAPK inhibitor SB202190 (Sigma Aldrich)  were dissolved in 0.1% bovine serum albumin or in dimethyl sulfoxide (DMSO), respectively. TNF and SB202190 were added directly to growth media to reach final concentrations of 10 ng/μl (TNF) and 100 μM (SB202190). Cells were collected at time points indicated in the results.
Total RNA was extracted from rMC-1 or primary Müller cells using the Megamax RNA isolation kit (Life Technologies) according to the manufacturer’s instructions. cDNA was prepared using the high capacity cDNA reverse transcription kit (Life Technologies). RT-PCR reactions were conducted using appropriate primer pairs (Additional file 8: Table S1). All primer pairs were checked for their amplification efficiency using serial dilutions of template and for the generation of a single amplicon of the correct size. Actb was used as internal control. Additional internal controls, Gapdh and Rpl32, were used for each new treatment. RT-PCR reactions were performed in a StepOne Real-Time PCR system with Fast SybrGreen master mix (Life Technologies) or a LightCycler 480 instrument with SybrGreen I Master mix (Roche, Basel, Switzerland). The comparative cycle threshold method was used to calculate relative transcript levels. Raw PCR data are presented in Additional file 9: File S3. N-values reflect independent experiments.
Primary mouse Müller cells (Müller cell enriched primary retinal cell culture)
Animal experimentation protocols were accepted by the Veterinary Authorities of Zurich and experiments adhered to the statement of ‘The Association for Research in Vision and Ophthalmology’ for the use of animals in research. Ttp −/− mice  and Rlbp-GFP transgenic mice  expressing GFP specifically in Müller glia cells were generously provided by Dr. Thomas Rülicke (University of Veterinary Medicine Vienna, Austria) and Dr. Edward M. Levine (University of Utah, Salt Lake City, UT, USA), respectively. Rlbp-GFP;Ttp −/− and Rlbp-GFP;Ttp +/− pups were euthanized by a CO2 overdose and decapitated between P8 and P12, and retinas were isolated. Retinal cells were dissociated according to the protocol by Siegert and colleagues . Dissociated cells were centrifuged for five minutes at 50 × g to increase the relative ratio of large GFP-positive cells. Cells from two retinas of individual mice were subdivided into eight wells for further treatments. Retinal cells in each well were cultured in 2 ml media for two weeks with a media change every two days using the same conditions as for the rMC-1 cells except that 25 mM D-sorbitol (Sigma-Aldrich) instead of glucose was used for Müller cell enrichment . Unlike adult Müller cells, GFP expression in P8-12 Müller cells was weak and upon attachment, GFP expression declined further, a phenomenon possibly due to proliferation of primary Müller cells or loss of cellular connections and increased surface area in culture. Müller cells proliferated and reached confluence generally after about two weeks.
Sequence alignment and ARE identification
Alignment of 3’UTR sequences from Lif genes of various mammals and identified highly conserved regions were retrieved from Vista pre-computed whole-genome alignments [56,57]. The mouse Lif 3′UTR sequence (NM_008501.2) was manually scanned for AU-rich regions according to Hao et al. and Caput et al. [50,109]. To qualify as an ARE, AUUU sequences needed to be accompanied by at least three additional A or U residues. In cases where several AUUU sequences were spaced by less than 3 bps, we checked for the presence of an AUUUA core motif. Putative micro RNA binding sites were identified using Targetscan software .
Cloning and site directed mutagenesis
PGL4.12[luc2CP] vector (Promega, Madison, WI, USA) was restriction digested with HindIII and XbaI to isolate luc2CP. The isolated luc2CP fragment (containing hCP1 and hPEST) was cloned into pGL3 control vector (Promega,) containing SV40 promoter and enhancer for robust expression, as Luc2CP luminescence was barely detectable in normal expression vectors due to the highly unstable nature of the protein. The resulting vector (ΔUTR) had a minimal 3′UTR and was used for further cloning. Mouse whole Lif 3′UTR (source: BAC clone RP23-451O6 (Children’s Hospital Oakland Research Institute, Oakland, CA, USA)), AU-rich region I, human β-Actin (ACTB) 3′UTR and human genomic sequences (alternate assembly CHM1_1.1, chr2:122988136–122991277) were PCR amplified and cloned into ΔUTR vector using the XbaI restriction site. Similarly, individual mouse Lif 3′UTR fragments were PCR amplified and cloned into the ΔUTR vector using XbaI and PstI restriction sites with the exception of AU-rich region II and region 27. These two regions were generated by restriction digestion of the plasmid containing the whole Lif 3′UTR using PstI-XcmI and PstI-EcoNI enzyme combinations for AU-rich region II and region 27, respectively. Overhanging ends of digested plasmids were blunted using Klenow enzyme and religated.
Site-directed mutagenesis was done according to instructions provided by the QuikChange Lightning Multi Site-Directed Mutagenesis kit (Agilent, Santa Clara, CA, USA). ARE core sequences were replaced by restriction digestion sites for identification of clones. During the mutation of element B (plasmid 27 ‘2 ARE’) within region 27, multiple ARE containing plasmids were generated as byproducts and used to test for the effects of increased numbers of ARE elements. Plasmid ‘27-B’ was generated by restriction digestion of plasmid ‘27-No’ with SacII and SpeI that was generated during mutation of elements A and C. Element B was introduced back to digested plasmid ‘27-No’ by annealed primers that contained element B and appropriate overhangs. Primers used for cloning are listed in Additional file 10: Table S2.
rMC-1 cells were transfected with constructs containing 3′UTR sequences fused to Luc2CP firefly luciferase as described previously . Renilla luciferase expressing vector, pRL-CMV (Promega), was used as internal control. Transfected rMC-1 cells were cultured for 24 hours and luciferase levels were measured using the Dual-Luciferase kit (Promega). Firefly/renilla luciferase ratios were calculated and expressed relative to the respective control. Each construct was tested in triplicate in three to four independent experiments.
In vitro transcription and 3′ biotin labeling
RNAs used for capturing RNA binding proteins from cell extracts were generated by in vitro transcription (Maxiscript kit; Life Technologies) using PCR amplified templates from region 27, region 36 and their respective ARE-null counterparts. Primers used for amplification of templates are listed in Additional file 10: Table S2. RNA probes were labeled with biotin at the 3′ end using the RNA 3′ End Biotinylation Kit (Pierce, Rockford, IL, USA). All procedures were followed according to the instructions from the manufacturers.
Quantitative mass spectrometry analysis and GO-pathway analysis
rMC-1 cells were grown in SILAC DMEM (GE Healthcare Life Sciences, Pittsburgh, PA, USA) supplemented with 3 mM L-glutamine (GE Healthcare Life Sciences), 10% dialyzed fetal bovine serum (GE Healthcare Life Sciences) and 0.55 mM lysine, 0.4 mM arginine. Light SILAC medium was supplemented with 12C6, 14N2 lysine and 12C6, 14N4 arginine. Heavy SILAC medium was supplemented with either 13C6 lysine and 13C6, 15N4 arginine or 13C6, 15N2 lysine and 13C6, 15N4 arginine. A total of 0.5mM proline was added to all SILAC media to prevent arginine to proline conversion. All amino acids were purchased from Silantes (Munich, Germany).
Biotin-labeled RNA (2 μg) was bound to Strep-tactin beads (IBA, Goettingen, Germany) in RNA binding buffer (150 mM NaCl, 50 mM Hepes-HCl pH 7.5, 0.5% NP40 (v/v), 10 mM MgCl2, Phosphatase Inhibitor Cocktail 2 and 3 (Sigma-Aldrich)) and incubated on a rotation wheel at 4°C. Beads were washed three times with RNA wash buffer containing 150 mM NaCl, 50 mM Hepes-HCl pH 7.5, 0.1% NP40 and 10 mM MgCl2 and Phosphatase Inhibitor Cocktail 2 and 3 (Sigma-Aldrich) before incubation at 4°C for 30 minutes with 2 mg of cytoplasmic extract, 200 units RNase inhibitor (Fermentas, Schwerte, Germany) and 20 μg yeast tRNA. After incubation, the corresponding samples were combined and the beads were washed another three times with RNA wash buffer before the protein/RNA complexes were eluted from the beads with Laemmli buffer. The eluted proteins were subjected to gel-based pre-fractionation and tryptic cleavage as described elsewhere [110,111].
Liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis was performed on an Ultimate3000 nano RSLC system (Thermo Fisher Scientific, Waltham, MA USA) coupled to a LTQ Orbitrap Velos mass spectrometer (Thermo Fisher Scientific) by a nano spray ion source. Tryptic peptide mixtures were automatically injected and separated by a linear gradient from 5% to 40% of buffer B (2% acetonitrile, 0.1% formic acid in HPLC grade water) in buffer A (0.1% formic acid in HPLC grade water) at a flow rate of 300 nl/minute over 90 minutes. Remaining peptides were eluted by a short gradient from 40% to 100% buffer B in five minutes. The eluted peptides were analyzed by the LTQ Orbitrap Velos mass spectrometer. From the high resolution MS pre-scan with a mass range of 300 to 1,500, the ten most intense peptide ions were selected for fragment analysis in the linear ion trap if they exceeded an intensity of at least 500 counts and if they were at least doubly charged. The normalized collision energy for CID was set to a value of 35 and the resulting fragments were detected with normal resolution in the linear ion trap. The lock mass option was activated, the background signal with a mass of 445.12002 was used as lock mass 5. Every ion selected for fragmentation was excluded for 20 seconds by dynamic exclusion.
All acquired spectra were processed and analyzed using the MaxQuant software 6 (version 18.104.22.168) and the human specific IPI database version 3.52  in combination with Mascot (Matrix Science, version 2.2). Cysteine carbamidomethylation was selected as fixed modification, and methionine oxidation and protein acetylation were allowed as variable modifications. The peptide and protein false discovery rates were set to 1%. Contaminants, such as keratins, were removed. Proteins, identified and quantified by at least two unique peptides were considered for further analysis. The significance values were determined by Perseus tool (part of MaxQuant) using significance A . GO pathway analyses were done using the web-based Gene Set Analysis Toolkit [113,114].
rMC-1 cells were seeded on six-well plates (50,000 cells per well, 2 ml growth medium). After 24 hours at 37°C and 5% CO2, rMC-1 cells were transfected with siRNA using RNAiMax (Life Technologies) and 80 pmol of specific siRNA oligonucleotides (Additional file 11: Table S3; Qiagen, Hilden, Germany) or AllStars Negative Control siRNA (Qiagen) according to the manufacturer’s instructions. Cells were used for experiments 48 hours after transfection.
rMC-1 cells were lysed in 200 μl 2 × Laemmli sample buffer. A total of 30 μl of the homogenate was separated on 10% SDS-polyacrylamide gels, blotted and probed as described previously . Primary antibodies for ELAVL1 (1:2,000, cat# sc-5261, Santa Cruz, Dallas, TX, USA) and ILF3 (1:1,000, cat# 19887-1-AP, Proteintech, Manchester, UK) were applied over night at 4°C. The secondary antibody (1:10,000, peroxidase-linked anti-rabbit immunoglobulin G (IgG), cat# NA934; GE Healthcare) was applied for one hour at room temperature. We have used WesternBright Sirius horseradish peroxidase (HRP) substrate (Advansta, Menlo Park, CA, USA) for chemiluminescence reaction. Fusion FX7 Advance imaging system (Vilber Lourmat, Torcy, France) with a CCD camera was used for digital signal detection. Recordings were taken at the dynamic range of exposure without binning. Calculations for expression levels were performed using BioD1 software (Vilber Lourmat) without background subtractions. Signals for ACTB served as controls.
For Western blotting experiments on SD, H2O2, SD + H2O2, TNF or SB treated rMC-1 cells (Additional file 2: Figure S2), the same procedures were applied except for the detection system and quantification method. Briefly, X-ray film-based detection was followed by Image J quantification relative to ACTB or unphosphorylated p38 MAPK levels. The following primary antibodies and dilutions were used: p38 MAPK (cat# 9212, 1:1,000, Cell Signaling, Danvers, MA, USA), phospho-p38 MAPK (cat# 9211, 1:1,000, Cell Signaling); phospho-HSP27 (cat# 2401P, 1:1,000, Cell Signaling); phospho-MKK3/6 (cat# 9231S, 1:1,000, Cell Signaling); ACTB (cat# A5441, 1:5,000, Sigma-Aldrich). Peroxidase-linked anti-mouse IgG (cat# sc-2031, Santa Cruz) was used at a dilution of 1:10,000 as secondary antibody.
129S6 wild type mice were or were not exposed to two hours of 15,000 lux of white light as described . Twenty-four hours after light exposure, mice were euthanized, eyes enucleated and fixed in 4% paraformaldehyde (PFA) prepared in phosphate buffered saline (PBS; pH 7.4), as described previously . Cryosections (12 μm) were blocked for one hour with 3% normal goat serum (containing 0.3% Triton X-100 in PBS), and incubated overnight at 4°C with rabbit anti-ILF3 (1:100; cat# 19887-1-AP, Proteintech, Manchester, UK), rabbit anti-KHSRP (1:250; cat# NBP1-18910, Novus Biologicals, Cambridge, UK) or mouse anti-glutamine synthetase (1:500; cat# MAB302, Millipore, Darmstadt, Germany) primary antibodies. Slides were washed three times with PBS and incubated with Cy2-labeled secondary anti-rabbit or Cy3-labeled secondary anti-mouse antibodies (Jackson ImmunoResearch Laboratories, Soham, UK), counterstained with 4',6-diamidino-2-phenylindole (DAPI) and analyzed by fluorescence microscopy (Axioplan 2; Carl Zeiss AG, Feldbach, Switzerland).
Statistical analysis was performed using ANOVA with appropriate posttests (see Figure legends) for multiple comparisons. Student's t-tests were used for individual pairwise comparisons. P values less than 0.05 were considered to indicate significant differences. Error bars represent the standard error of the mean (SEM). Graph Pad 6 software (GraphPad Inc., San Diego, USA) was used for all statistical analyses.
We thank Dr. Vijay Sarthy (Northwestern University, Chicago, IL) for providing rMC-1 cells. We also thank Dr. Thomas Rülicke (University of Veterinary Medicine Vienna, Austria) for the TTP knockout mouse line, Dr. Edward M. Levine (University of Utah, Salt Lake City, UT) for the Rlbp-GFP transgenic mouse line, Dr. Sandrine Joly (University of Zurich) for her help with initial cell culture experiments and Nicola Horn for excellent technical assistance. This work was supported by the Swiss National Science Foundation (31003A_133043 and 31003A_149311 to CG), the Velux Foundation (to CA), the European Community’s Seventh Framework Program FP7 (grant agreement no. 241955; SYSCILIA to MU and grant agreement no. 278568; PRIMES to MU and KB) and the Kerstan Foundation (to MU). Funding for open access charge: Swiss National Science Foundation.
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