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Open Access

Expression of leukemia inhibitory factor in Müller glia cells is regulated by a redox-dependent mRNA stability mechanism

  • Cavit Agca1, 6,
  • Karsten Boldt2,
  • Andrea Gubler1,
  • Isabelle Meneau1,
  • Armelle Corpet3, 7,
  • Marijana Samardzija1,
  • Manuel Stucki3,
  • Marius Ueffing2 and
  • Christian Grimm1, 4, 5Email author
BMC Biology201513:30

https://doi.org/10.1186/s12915-015-0137-1

Received: 7 January 2015

Accepted: 8 April 2015

Published: 25 April 2015

Abstract

Background

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.

Results

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.

Conclusions

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.

Keywords

LIFRedox signalingmRNA stabilityRetinaMüller glial cellsILF3KHSRPp38 MAPKNeuroprotection

Background

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 [2] 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 [20], 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 [39] and to induce axonal regeneration in zebrafish [40], 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 [21], 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.

Results

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].

Since LIF is one of the critical factors for photoreceptor survival and is expressed in Müller cells in response to photoreceptor injury [3,13,16,44], we tested the effect of subtoxic doses of H2O2 [45-47] on Lif mRNA expression in rMC-1 Müller cells [48]. Since rMC-1 cells expressed high levels of Lif mRNA under normal (optimal) culture conditions, stress-related signaling molecules like H2O2 were without effect and failed to super induce Lif expression (Additional file 1: Figure S1). As Müller glia may experience suboptimal (stressed) conditions in an injured retina, we serum deprived (SD) cells to impose a mild stress during the period of H2O2 signaling in vitro. This caused a rapid and strong reduction of Lif mRNA levels in Müller cells by 87% within one and 91% within two hours (Figure 1A). Whether this was due to the suboptimal conditions, or to the absence of Lif-inducing factors potentially present in serum has not been addressed. Importantly, SD allowed us to investigate the effect of H2O2 on Lif levels. Addition of H2O2 partially prevented this downregulation and resulted in Lif transcript levels that were 3.6-fold (one hour, P <0.001) and 4.5-fold (two hours, P <0.01) higher than in cells treated with SD alone (Figure 1A). H2O2 needed to be present from the beginning of SD treatment since its late addition (after one hour of SD treatment) did not affect Lif levels (Figure 1A). Similarly, H2O2 had no effect on Lif expression when added to rMC-1 cells under normal cell culture conditions (Additional file 1: Figure S1). The effect of H2O2 was specific for Lif since the mRNA levels of other genes known to be expressed in Müller cells, such as ciliary neurotrophic factor (Cntf), Fgf2 and Gfap [2,13,49], or known to be postranscriptionally regulated at the level of mRNA stability, such as Ttp [42,50], were not increased by the addition of H2O2. Rather, Cntf was downregulated by 18% (one hour, P <0.01) and 27% (two hours, P <0.0001) in SD cells treated with H2O2 (Figure 1A).
Figure 1

H2O2 stabilizes Lif transcripts in rMC-1 Müller cells. A) rMC-1 cells were serum deprived (SD, circle) for two hours. H2O2 (50 μM) was added immediately (SD + H2O2, square) or at one hour during SD (SD + late H2O2, triangle). B) rMC-1 cells were treated with actinomycin D (ActD, circle) for two hours. H2O2 was added immediately (ActD + H2O2, square) or at one hour during ActD treatment (ActD + late H2O2, triangle). RNA levels of the indicated genes were determined by real-time PCR at 0 hour, 1 hour and 2 hours of treatment and expressed relative to levels at 0 hour. Shown are means ± SEM of N = 4 to 5. One-way ANOVA with Sidak’s posttests were used to compare RNA levels against levels after SD (A) or ActD (B) treatments at one hour and two hours. (*) P <0.05, (**) P <0.01, (***) P <0.001 and (****) P <0.0001. ANOVA, analysis of variance; SEM, standard error of the mean.

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.

Since it was shown that retinal degenerations activate Lif mRNA transcription [21,50] and H2O2 clearly affected a posttranscriptional process, we tested whether H2O2 can increase longevity of Lif mRNA after an initial boost of mRNA production. We treated rMC-1 cells with TNF, which was previously shown to transiently upregulate both Lif and Ttp transcription [21,50]. When applied at one hour of TNF-induced upregulation of gene transcription, SD and ActD treatment caused a rapid decline of Lif and Ttp mRNA levels. This decline was partially prevented by the addition of H2O2 suggesting that redox signaling can also stabilize Lif and Ttp mRNAs under conditions of increased gene activity (Figure 2A, B). The data argue that H2O2 may function as a signaling molecule to overcome the unstable nature of Lif and Ttp mRNAs resulting in a more sustained expression of these genes when needed.
Figure 2

H2O2 stabilizes Lif transcripts after TNF induced gene expression in rMC-1 Müller cells. rMC-1 cells were treated with tumor necrosis factor-alpha (TNF, circle) for one hour to induce Lif gene expression. Following TNF treatment cells were either (A) serum deprived (SD, square) or (B) incubated with ActD (square). H2O2 (50 μM) was added either at the one hour (+ H2O2, triangle) or the two hour time-point (+ late H2O2, del). RNA levels of Lif and Ttp were determined by real-time PCR at the indicated time-points and expressed relative to levels at 0 hour. Shown are means ± SEM of N = 4. One-way ANOVA with Sidak’s posttests were used to compare RNA levels against levels after SD or ActD treatments at two hours and three hours. (*) P <0.05, (**) P <0.01, (***) P <0.001. ActD, actinomycin D. ANOVA, analysis of variance; SEM, standard error of the mean.

As already reported earlier, inhibition of p38 MAPK by SB202190 (SB) strongly decreased Lif mRNA levels in rMC-1 cells (Figure 3A, [21]). Similar to the effect after SD or ActD treatment, addition of H2O2 to SB-treated cells increased Lif mRNA levels. This argues that redox dependent Lif mRNA stabilization may not depend on p38 MAPK, even though several components of its signaling system are activated by H2O2 (Additional file 2: Figure S2A, B) [51-53]. To strengthen this conclusion, we tested whether H2O2 might be able to increase Lif mRNA levels in the absence of TTP, an important component of the p38 MAPK-dependent mRNA stability pathway [41-43]. Indeed, H2O2 rescued the SD-induced downregulation of Lif mRNA levels also in primary Müller cells isolated from Ttp −/− mice (Figure 3B). Importantly, these experiments also showed that redox regulation of Lif was not specific for rMC-1 cells, but was also a feature of primary Müller cells making it likely that this mechanism is relevant for the regulation of Lif expression in vivo.
Figure 3

H2O2-induced stabilization of Lif in rMC-1 and primary Müller cells is independent of p38 MAPK. A) rMC-1 cells were (+) or were not (−) treated with p38 MAPK inhibitor SB202190 (SB) in the presence (+) or absence (−) of H2O2 (50 μM) for one hour. RNA levels were determined by real-time PCR and expressed relative to the levels in untreated cells. Shown are means ± SEM of N = 6 (five independent experiments). One-way ANOVA with Tukey’s posttests were used to compare all pairs of columns. B) Primary mouse Müller cells isolated from Rlbp-GFP mice heterozygous (Ttp +/− ) or homozygous (Ttp −/− ) for the Ttp knockout allele were (+) or were not (−) serum deprived (SD) for two hours in the presence (+) or absence (−) of H2O2 (50 μM). Lif mRNA levels were determined by real-time PCR at two hours of treatment and expressed relative to the levels of untreated cells. Shown are means ± SEM of N = 3 to 4. One-way ANOVA with Sidak’s posttests were used to compare the effect of treatment for both Ttp +/− and Ttp −/− Müller cells. (**) P <0.01 and (***) P <0.001. ANOVA, analysis of variance; SEM, standard error of the mean.

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

Since Lif encodes a highly unstable mRNA (Figures 1 and 2) and stability of particular RNAs like Cox2 can be regulated through proteins that bind to AREs within their 3′UTRs [54,55], we analyzed the sequence of the 3,180 bp long Lif 3′UTR for the presence of AREs. Alignment of the 3′UTRs from various mammalian Lif genes [56,57] showed that several regions were strongly conserved (Figure 4A). Two large regions (termed AU-rich R I and AU-rich R II) were identified that were particularly rich in adenosine and uracil (Figure 4A). These two regions contained several smaller regions with AUUU sequence elements and AUUUA core motifs (regions 27, 31, 36). Region 35 was an exception in that it contained only a string of 7 AUUU sequences without core motif. In addition, two conserved putative binding sites for miR-29 (region 36) and miR-17 (region 38) were identified using TargetScan software [58]. No AREs or miRNA binding sites were detected in regions 32 and 37 (Figure 4A, Additional file 3: Figure S3A).
Figure 4

The Lif 3′UTR contains conserved regions with AREs that influence RNA levels in rMC-1 cells. A) Sequence alignment of the 3,180 bp long Lif 3′UTR sequences from Mus musculus (Mm), Homo sapiens (Hs), Rhesus monkey (Rm), Canis lupus familiaris (Cf), Bos taurus (Bt) and Pan troglodytes (Pt) revealed two highly conserved AU-rich regions (R I and R II) and seven smaller regions containing AREs (R27, R31, R32, R35, R37), a miRNA binding site (R38) or both (R36). B, C) rMC-1 cells were transfected with luciferase-reporter constructs, as indicated. Luminescence levels were determined after 24 hours and expressed relative to a construct with a minimal 3′UTR (ΔUTR). D) rMC-1 cells were transfected with luciferase-reporter constructs carrying R36, R27, R32 or ΔUTR sequences. Gene expression was blocked with ActD 24 hours after transfection. Luminescence levels were determined at 0 to 4 hours after ActD addition and shown relative to the levels of the ΔUTR control construct that were set to 1 for each time point. Shown are means ± SEM of N = 4 to 5. One-way ANOVA (B, C) or two-way ANOVA (D) with Dunnett’s posttests were used to compare constructs to the levels of the ΔUTR construct. (*) P <0.05, (**) P <0.01, (***) P <0.001 and (****) P <0.0001. ANOVA, analysis of variance; AREs, AU-rich elements; SEM, standard error of the mean.

To evaluate the potential importance of these conserved elements for the regulation of Lif RNA stability, we fused fragments of the mouse 3′UTR to a luciferase gene driven by the SV40 promoter and enhancer (Table 1, Additional file 3: Figure S3B). The incorporation of CL1 [59] and PEST [60] sequences led to the production of an unstable luciferase protein (Luc2CP) suitable to monitor changes in mRNA levels by measuring luminescence in cell culture homogenates.
Table 1

Vector list of cloned regions used in constructs

Vector name

Lif 3′UTR position (1 to 3180)

Length of 3′UTR (bp)

AREs

Sequence source

Cloning method

∆UTR

0

0

0

NA

RE

Whole Lif 3′UTR

1 to 3180

3180

11

mus musculus

PCR/RE

hgen DNA

NA

3142

2

homo sapiens

PCR/RE

AU-rich R I

1 to 570

570

4

mus musculus

PCR/RE

AU-rich R II

2556 to 3180

625

5

mus musculus

RE

hACTB 3′UTR

NA

600

3

homo sapiens

PCR/RE

R27 (wt)

2999 to 3180

182

3

mus musculus

RE

R32

2805 to 2998

194

0

mus musculus

PCR/RE

R31

2541 to 2812

272

2

mus musculus

PCR/RE

R38

2079 to 2245

167

0

mus musculus

PCR/RE

R36 (wt)

1062 to 1231

170

2

mus musculus

PCR/RE

R35

398 to 570

173

4

mus musculus

PCR/RE

R37

62 to 229

168

0

mus musculus

PCR/RE

R27-A

2999 to 3180

182

1

mus musculus

SD

R27-B

2999 to 3180

182

1

mus musculus

RE/AN

R27-C

2999 to 3180

182

1

mus musculus

SD

R27-No

2999 to 3180

182

0

mus musculus

SD

R36-DE

1062 to 1231

170

2

mus musculus

SD

R36-D

1062 to 1231

170

1

mus musculus

SD

R36-E

1062 to 1231

170

1

mus musculus

SD

R36-No

1062 to 1231

170

0

mus musculus

SD

R27 2 ARE

2999 to 3180

182

2

mus musculus

SD

R27 4 ARE

2999 to 3180

237

4

mus musculus

SD

R27 9 ARE

2999 to 3180

351

9

mus musculus

SD

Name and details of position, length, number of putative AREs, and cloning strategies are listed for individual 3′UTR sequences used in the luciferase experiments. AREs, AU-rich elements; hgen, human genomic DNA; NA, not applicable; PCR, polymerase chain reaction; RE, restriction digestion; RE/AN, restriction digestion/primer annealing; SD, site-directed mutagenesis.

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.

In order to identify the functional elements within regions 27 and 36, we mutated individual AUUU sequences (Figure 5, Table 1, Additional file 4: Figure S4). Region 27 had three predicted AREs (A, B and C) that, when mutated, significantly increased reporter expression (Figure 5A). Mutating two elements in either combination resulted in Luc2PC levels similar to the triple mutant suggesting that none of the individual AREs was sufficient to significantly reduce Luc2PC expression but that the elements may need to interact or to act synergistically to regulate mRNA stability. Consistent with a synergistic effect, increasing the number of AREs within the same reporter construct gradually decreased reporter expression to levels found for the whole Lif 3′UTR (Figure 5B, Additional file 4: Figure S4B). Region 36 contained two predicted AREs (D and E) and a conserved miR-29 binding site (Additional file 3: Figure S3A). Mutating the miR-29 binding sequence alone or in combination with either ARE did not significantly alter Luc2CP levels. In contrast, simultaneous elimination of all three elements significantly increased Luc2CP levels (Figure 5C).
Figure 5

AREs are involved in cis-regulation of Lif mRNA stability in rMC-1 Müller cells. A) rMC-1 cells were transfected with reporter constructs fused to wild type or mutated R27 sequences (see Additional file 4: Figure S4A), as indicated. B) rMC-1 cells were transfected with reporter constructs fused to R27 sequences containing increasing numbers of AREs (see Additional file 4: Figure S4B), as indicated. C) rMC-1 cells were transfected with reporter constructs fused to indicated wild type or mutated R36 sequences (see Additional file 4: Figure S4C), as indicated. Luminescence levels were always determined after 24 hours and shown relative to the level generated by the ΔUTR control construct. Shown are means ± SEM of N = 3 to 4. Squares in horizontal bars indicate AREs or an miR-29 binding site, respectively, in the analyzed sequences. One-way ANOVA with Dunnett’s posttests were used to compare expression levels to constructs containing the respective wild type sequence (A, C) or to ‘2 ARE’ (B). (*) P <0.05, (**) P <0.01, (***) P <0.001 and (****) P <0.0001. ANOVA, analysis of variance; AREs, AU-rich elements; SEM, standard error of the mean.

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

To identify proteins that specifically bind to AREs in regions 27 and 36 we incubated the respective RNA sequences or their ARE-null counterparts (Additional file 4: Figure S4A, C) with protein extracts from rMC-1 cells. Mass spectrometry (MS)-based quantitative proteomics identified those proteins that preferentially bound to the wild type sequences of regions 27 or 36 (Tables 2 and 3; the mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium [61] via the PRIDE partner repository with the dataset identifier PXD001463). Gene Ontology (GO)-pathway analysis on identified proteins showed that pools were enriched in RNA binding proteins and contained 11 (region 27) and 14 (region 36) such proteins (Additional file 5: Figure S5A, B; blue circles, Additional file 6: File S1, Additional file 7: File S2). Whereas RNA binding proteins identified with region 36 were mainly linked to translational initiation and elongation as well as RNA catabolic processes (Table 3, stars, Additional file 7: File S2), region 27 RNA binding proteins were primarily associated with double-stranded RNA binding and mRNA stability (Table 2, stars, Additional file 6: File S1). Interestingly, the protein pool identified with region 27 was also enriched with cell redox homeostasis proteins (Table 2, squares; Additional file 5: Figure S5A, blue square, Additional file 6: File S1).
Table 2

List of RNA binding proteins associated with region 27 of the Lif 3′UTR

Protein ID

 

Protein name

Gene name

27 wt versus 27-No Significance

27 wt versus 27-No Rel. levels

Q15717

ELAV-like protein 1

ELAVL1

1.88E-08

8.50

P13693

 

Translationally-controlled tumor protein

TPT1

0.0005

3.69

Q01469

 

Fatty acid-binding protein, epidermal

FABP5

0.0007

3.58

Q12906

Interleukin enhancer-binding factor 3

ILF3

0.0023

3.11

P07741

 

Adenine phosphoribosyltransferase

APRT

0.0065

2.74

O75390

 

Citrate synthase, mitochondrial

CS

0.0069

2.72

Q9Y281

 

Cofilin-2

CFL2

0.0088

2.62

Q15185

 

Prostaglandin E synthase 3

PTGES3

0.0091

2.62

Q04760

 

Lactoylglutathione lyase

GLO1

0.0095

2.60

A6NL28

 

Putative tropomyosin alpha-3 chain-like protein

PTA3CLP

0.0103

2.57

Q12905

Interleukin enhancer-binding factor 2

ILF2

0.0119

2.52

P61956

 

Small ubiquitin-related modifier 2-3-4

SUMO2-3-4

0.0131

2.48

P48735

 

Isocitrate dehydrogenase [NADP], mitochondrial

IDH2

0.0150

2.43

Q96SI9

Spermatid perinuclear RNA-binding protein

STRBP

0.0155

2.42

P62633

 

Cellular nucleic acid-binding protein

CNBP

0.0173

2.38

P09104

 

Gamma-enolase

ENO2

0.0200

2.33

P30086

 

Phosphatidylethanolamine-binding protein 1

PEBP1

0.0214

2.31

P20962

 

Parathymosin

PTMS

0.0219

2.30

P61326

Protein mago nashi homolog-B

MAGOH-B

0.0225

2.29

P50395

 

Rab GDP dissociation inhibitor beta

GDI2

0.0233

2.28

Q14019

 

Coactosin-like protein

COTL1

0.0234

2.27

P07737

 

Profilin-1

PFN1

0.0242

2.26

P00505

 

Aspartate aminotransferase, mitochondrial

GOT2

0.0243

2.26

Q9BTT0

 

Acidic leucine-rich nuclear phosphoprotein 32 family member E

ANP32E

0.0248

2.25

O00299

 

Chloride intracellular channel protein 1

CLIC1

0.0268

2.22

Q9NR31

 

GTP-binding protein SAR1a

SAR1A

0.0280

2.21

Q8NBS9

Thioredoxin domain-containing protein 5

TXNDC5

0.0289

2.20

P29558

RNA-binding motif, single-stranded-interacting protein 1

RBMS1

0.0311

2.17

Q99471

 

Prefoldin subunit 5

PFDN5

0.0330

2.15

P13667

Protein disulfide-isomerase A4

PDIA4

0.0336

2.14

P60903

 

Protein S100-A10

S100A10

0.0365

2.11

P61088

 

Ubiquitin-conjugating enzyme E2 N

UBE2N

0.0381

2.10

P09382

 

Galectin-1

LGALS1

0.0410

2.07

P62834

 

Ras-related protein Rap-1A

RAP1A

0.0428

2.05

P09972

 

Fructose-bisphosphate aldolase C

ALDOC

0.0435

2.05

P30040

 

Endoplasmic reticulum resident protein 29

ERP29

0.0440

2.04

O60493

 

Sorting nexin-3

SNX3

0.0451

2.04

P22626

Heterogeneous nuclear ribonucleoproteins A2/B1

HNRNPA2B1

0.0456

2.03

Q15843

 

NEDD8

NEDD8

0.0464

2.03

P30101

Protein disulfide-isomerase A3

PDIA3

0.0464

2.03

P19338

Nucleolin

NCL

0.0470

2.02

O15511

 

Actin-related protein 2/3 complex subunit 5

ARPC5

0.0481

2.01

P00338

 

L-lactate dehydrogenase A chain

LDHA

0.0501

1.20

Q15121

 

Astrocytic phosphoprotein PEA-15

PEA15

0.0517

1.99

P62937

 

Peptidyl-prolyl cis-trans isomerase A

PPIA

0.0556

1.96

P14174

 

Macrophage migration inhibitory factor

MIF

0.0559

1.96

Q14103

Heterogeneous nuclear ribonucleoprotein D0

HNRNPD

0.0559

1.96

Q99729

Heterogeneous nuclear ribonucleoprotein A/B

HNRNPAB

0.0587

1.94

O14979

Heterogeneous nuclear ribonucleoprotein D-like

HNRPDL

0.0602

1.93

P10599

Thioredoxin

TXN

0.0602

1.93

RNAs of region 27 (‘R27 wt’) and its counterpart ARE-null region 27 (‘R27-No’) were used for RNA binding assays. Bound proteins were identified by quantitative mass spectrometry analysis and relative ratios were determined using total sequence reads for region ‘R27 wt’ and region ‘R27-No’. RNA binding proteins, (stars) and cell redox homeostasis proteins (squares) are indicated as identified by GO pathway analysis,(Additional file 6: File S1). The significance is the significance A P-value as determined by the Perseus software, defining the likelihood of a protein being enriched compared to the background distribution [112].

Table 3

List of RNA binding proteins associated with region 36 of the Lif 3′UTR

Protein ID

 

Protein name

Gene name

36 wt versus 36-No Significance

36 wt versus 36-No Rel. levels

Q13126

 

S-methyl-5-thioadenosine phosphorylase

MTAP

6.70E-06

2.11

Q9Y3U8

 

60S ribosomal protein L36

RPL36

1.43E-05

2.05

P30040

 

Endoplasmic reticulum resident protein 29

ERP29

8.03E-05

1.91

P62942

 

Peptidyl-prolyl cis-trans isomerase FKBP1A

FKBP1A

0.0003

1.80

Q14019

 

Coactosin-like protein

COTL1

0.0005

1.76

O75390

 

Citrate synthase, mitochondrial

CS

0.0012

1.68

P15121

 

Aldose reductase

AKR1B1

0.0015

1.66

Q9Y281

 

Cofilin-2

CFL2

0.0019

1.64

Q02543

60S ribosomal protein L18a

RPL18A

0.0020

1.64

P30041

 

Peroxiredoxin-6

PRDX6

0.0022

1.63

P42766

60S ribosomal protein L35

RPL35

0.0024

1.62

Q96AG4

 

Leucine-rich repeat-containing protein 59

LRRC59

0.0041

1.57

Q01130

Serine/arginine-rich splicing factor 2

SRSF2

0.0051

1.55

P26447

 

Protein S100-A4

S100A4

0.0080

1.51

P60059

 

Protein transport protein Sec61 subunit gamma

SEC61G

0.0089

1.50

Q96AE4

Far upstream element-binding protein 1

FUBP1

0.0090

1.50

P09493

 

Tropomyosin alpha-1 chain

TPM1

0.0099

1.49

P40616

 

ADP-ribosylation factor-like protein 1

ARL1

0.0104

1.48

Q99536

 

Synaptic vesicle membrane protein VAT-1 homolog

VAT1

0.0115

1.47

P84098

60S ribosomal protein L19

RPL19

0.0120

1.47

P61019

 

Ras-related protein Rab-2A

RAB2A

0.0127

1.47

Q9HAV7

 

GrpE protein homolog 1, mitochondrial

GRPEL1

0.0127

1.47

O43396

 

Thioredoxin-like protein 1

TXNL1

0.0136

1.46

P26639

 

Threonine--tRNA ligase, cytoplasmic

TARS

0.0137

1.46

P09972

 

Fructose-bisphosphate aldolase C

ALDOC

0.0143

1.45

Q01581

 

Hydroxymethylglutaryl-CoA synthase, cytoplasmic

HMGCS1

0.0144

1.45

P63261

 

Actin, cytoplasmic 2; Actin, cytoplasmic 2, N-terminally processed

ACTG1

0.0152

1.45

Q99497

Protein DJ-1

PARK7

0.0180

1.43

P16989

DNA-binding protein A

CSDA

0.0192

1.42

P06748

Nucleophosmin

NPM1

0.0212

1.41

Q9BTT0

 

Acidic leucine-rich nuclear phosphoprotein 32 family member E

ANP32E

0.0221

1.41

P39023

60S ribosomal protein L3

RPL3

0.0227

1.41

P13693

 

Translationally-controlled tumor protein

TPT1

0.0267

1.39

P41567

Eukaryotic translation initiation factor 1

EIF1

0.0270

1.39

P49207

60S ribosomal protein L34

RPL34

0.0290

1.38

Q9Y266

 

Nuclear migration protein nudC

NUDC

0.0305

1.38

Q15020

Squamous cell carcinoma antigen recognized by T-cells 3

SART3

0.0331

1.37

P09429

 

High mobility group protein B1; Putative high mobility group protein B1-like 1

HMGB1

0.0364

1.36

P40429

 

60S ribosomal protein L13a; Putative 60S ribosomal protein L13a-like

RPL13A

0.0365

1.36

O43747

 

AP-1 complex subunit gamma-1

AP1G1

0.0400

1.35

Q00577

 

Transcriptional activator protein Pur-alpha

PURA

0.0410

1.35

P27635

 

60S ribosomal protein L10

RPL10

0.0414

1.34

P00558

 

Phosphoglycerate kinase 1

PGK1

0.0462

1.33

P22314

 

Ubiquitin-like modifier-activating enzyme 1

UBA1

0.0473

1.33

Q9H0U4

 

Ras-related protein Rab-1B

RAB1B

0.0475

1.33

P61106

Ras-related protein Rab-14

RAB14

0.0495

1.32

P49588

 

Alanine--tRNA ligase, cytoplasmic

AARS

0.0503

1.32

O75915

PRA1 family protein 3

ARL6IP5

0.0505

1.32

P55795

 

Heterogeneous nuclear ribonucleoprotein H2

HNRNPH2

0.0523

1.32

P30086

 

Phosphatidylethanolamine-binding protein 1; Hippocampal cholinergic neurostimulating peptide

PEBP1

0.0533

1.32

P68036

 

Ubiquitin-conjugating enzyme E2 L3

UBE2L3

0.0580

1.31

Q9UQE7

 

Structural maintenance of chromosomes protein 3

SMC3

0.0583

1.31

RNAs of region 36 (‘R36 wt’) and its counterpart ARE-null region 36 (‘R36-No’) were used for RNA binding assays. Bound proteins were identified by quantitative mass spectrometry analysis and relative ratios were determined using total sequence reads for region ‘R36 wt’ and region ‘R36-No’. RNA binding proteins (stars) are indicated as identified by GO pathway analysis (see Additional file 7: File S2). The significance is the significance A P-value as determined by the Perseus software, defining the likelihood of a protein being enriched compared to the background distribution.

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 [64]. ILF3 was additionally shown to interact with several RNA binding proteins including heterogeneous nuclear ribonucleoprotein (HNRNP) D, HNRNPA2/B1 and HNRNPA/B [66]. 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.

To test whether the identified proteins are indeed involved in redox regulation of Lif mRNA stability we silenced their expression in rMC-1 cells and analyzed the effect of H2O2 on Lif mRNA levels. Ilf3 silencing by two different siRNAs significantly impaired Lif mRNA stabilization in H2O2-treated serum-deprived cells by about 50%. Knockdown of Elavl1 or Hnrnpd, however, had no significant effect (Figure 6A). Controls showed that transfection of siRNA decreased expression levels of target mRNAs by 86% (Elavl1), 79% (Ilf3; si1), 52% (Ilf3; si2) and 91% (Hnrnpd), as compared to cells transfected with scrambled siRNA (Ctrl, Figure 6B). Similarly, protein levels of ELAVL1 and ILF3 (even in the case of Ilf3 siRNA2) were severely reduced in cells treated with the respective siRNAs (Figure 6B, lower panels). Silencing of either gene did not significantly alter Lif expression in untreated Müller cells (Figure 6C).
Figure 6

ILF3 regulates redox dependent mRNA stabilization in rMC-1 Müller cells. A) rMC-1 cells were transfected with siRNAs directed against Ilf3, Elavl1 or Hnrnpd, as indicated. Scrambled siRNA was used as control (Ctrl). Medium was changed 48 hours after transfection and then cells were (SD) or were not (C) serum deprived. H2O2 (50 μM) was added to the indicated samples. RNA levels were determined by real-time PCR two hours after treatment and expressed relative to untreated cells (C), which were set to 1 for each knockdown series. N = 4 to 8. B) Top panel: rMC-1 cells were transfected with siRNAs directed against Ilf3, Elavl1 or Hnrnpd, as indicated. Scrambled siRNA was used as control (Ctrl). Medium was changed 48 hours after transfection and respective target RNA levels were determined after two hours by real-time PCR. Bottom panels show protein levels at 48 hours after siRNA transfection. N = 4. C) rMC-1 cells were transfected with siRNAs directed against Ilf3, Elavl1 or Hnrnpd, as indicated. Scrambled siRNA was used as control (Ctrl). Medium was changed 48 hours after transfection and Lif RNA levels were determined after two hours by real-time PCR. To knockdown Ilf3 expression, two different siRNAs (Ilf3 (si1), Ilf3 (si2)) were used to exclude off-target effects. Shown are means ± SEM of N = 4. One-way ANOVA with Dunnett’s posttests was used to compare Lif levels after SD + H2O2 treatments (A) and in untreated knockdowns against control (C). Student’s t-test was used to compare downregulation of target genes (B). (**) P <0.01, (***) P <0.001. ANOVA, analysis of variance; ILF3, interleukin enhancer binding factor 3; SEM, standard error of the mean.

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

We identified ILF3 to be of importance for regulating Lif mRNA levels in redox signaling, probably through its interaction with AREs in the 3′UTR of the gene transcript. However, ILF3 did not affect Lif mRNA levels under normal growth conditions. We also showed that TTP, known to be capable of regulating the stability of ARE-containing mRNAs in various systems, was not involved in controlling Lif mRNA levels under normal conditions (Figure 3B). Thus, we asked whether other RNA binding proteins may affect Lif mRNA levels under normal non-stressed growth conditions. KHSRP was an obvious candidate as it is a multifunctional RNA binding protein involved in both transcriptional and posttranscriptional gene regulation as well as in miRNA biogenesis [70-73]. Furthermore, KHSRP is regulated through p38 MAPK signaling [71] that was shown to be important for Lif mRNA transcription [21]. Silencing Khsrp to 15% (si1) or 8% (si2) of control levels resulted in 78% (si1) and 52% (si2), respectively, lower Lif mRNA levels under normal conditions (Figure 7A, B) suggesting that KHSRP is essential to maintain normal Lif mRNA levels in non-stressed cells. However, lack of KHSRP did not affect redox-mediated stabilization of Lif transcripts. Note that expression levels of Lif in Figure 7C have been normalized to the respective controls (set to 1) to better visualize the effect of H2O2 in Khsrp knockdown cells. These results argue that KHSRP is involved in Lif expression independently of redox regulated mRNA stabilization and suggest that mechanisms controlling Lif mRNA levels under normal conditions differ from those under stress conditions. KHSRP may be involved in the first, ILF3 in the second, situation. Both proteins showed widespread expression in the normal mouse retina. Cells of the inner nuclear layer and of the ganglion cell layer were intensely stained whereas photoreceptor cells appeared to contain lower levels of the two proteins. Light-induced injury did not affect the expression pattern. Co-staining for glutamine synthetase confirmed expression of ILF3 and KHSRP in Müller glia supporting a role of ILF3 and KHSRP in Müller cells in vivo (Figure 8).
Figure 7

Silencing of Khsrp reduces Lif mRNA levels in non-stressed rMC-1 Müller cells. A, B) rMC-1 cells were transfected with siRNAs directed against Khsrp. Scrambled siRNA was used as control (Ctrl). Medium was changed 48 hours after transfection and RNA levels for Khsrp (A) or Lif (B) were determined after two hours by real-time PCR. C) rMC-1 cells were transfected with siRNAs directed against Khsrp. Scrambled siRNA was used as control (Ctrl). Medium was changed 48 hours after transfection and cells were (SD) or were not (C) serum deprived. H2O2 (50 μM) was added to samples as indicated. RNA levels were determined by real-time PCR two hours after treatment and expressed relative to untreated cells (C), which were set to 1 for each knockdown series. Two different siRNAs (Khsrp (si1), Khsrp (si2)) were used to exclude off-target effects. Shown are means ± SEM of N = 3 to 4. Student’s t-test was used to compare downregulation of Khsrp and Lif in untreated cells (A, B). One-way ANOVA with Dunnett’s posttests were used to compare Lif or Khsrp levels to controls (A, B) or Lif levels after SD + H2O2 treatments (C). (***) P <0.001. ANOVA, analysis of variance; SEM, standard error of the mean.

Figure 8

Immunofluorescence of ILF3 and KHSRP in the retina. Retinal sections were prepared from control mice and from mice at one day after light exposure and stained (A) for glutamine synthetase (GS, green) and ILF3 (red) or (B) for GS (green) and KHSRP (red). (C) Control stainings with secondary antibodies only (as indicated). DAPI was used to visualize nuclei. DAPI, 4',6-diamidino-2-phenylindole.

Discussion

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 [74]. 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 [21]. 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 [2]. 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 [81] and aquaporin-4 has been shown to alter its localization in Müller glia cells upon photoreceptor injury [82], 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 [8]. 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 [84]. 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 [50]. 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 [86], 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) [91], 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 [64]. 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 [64].

Interestingly, neither Lif nor Mkp1 mRNAs were identified as ILF3 target RNAs by a high-throughput ribonucleoprotein immunoprecipitation assay [94] even though both Lif (this study) and Mkp1 (Kuwano et al. [64]) were shown to be regulated by ILF3 during oxidative stress. Since non-treated HeLa cell lysates were used for those experiments [95], 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 [94], 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 [61]). 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 [73], 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 [97]. 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 [71] 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 [100] 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.

Conclusions

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.

Methods

Cell culture, serum deprivation, H2O2, ActD, TNF and p38 MAPK inhibitor treatment

The rat Müller glia cell line rMC-1 [48] 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 [21]. 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) [104] 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.

RT-PCR analysis

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 [105] and Rlbp-GFP transgenic mice [106] 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 [107]. 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 [108]. 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 [58].

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.

Luciferase assay

rMC-1 cells were transfected with constructs containing 3′UTR sequences fused to Luc2CP firefly luciferase as described previously [21]. 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 1.3.0.5) and the human specific IPI database version 3.52 [112] 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 [112]. GO pathway analyses were done using the web-based Gene Set Analysis Toolkit [113,114].

siRNA transfection

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.

Western blotting

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 [13]. 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.

Immunofluorescence

129S6 wild type mice were or were not exposed to two hours of 15,000 lux of white light as described [115]. 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 [116]. 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).

Data analysis

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.

Declarations

Acknowledgements

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.

Authors’ Affiliations

(1)
Department of Ophthalmology, Lab for Retinal Cell Biology, University of Zurich
(2)
Division of Experimental Ophthalmology and Medical Proteome Center, Centre for Ophthalmology, University of Tübingen
(3)
Department of Gynecology, University of Zurich
(4)
Zurich Center for Integrative Human Physiology (ZIHP), University of Zurich
(5)
Neuroscience Center (ZNZ), University of Zurich
(6)
Present address: Department of Biomedicine, University Hospital Basel
(7)
Present address: Center for Molecular and Cellular Physiology and Genetics, University Lyon I

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© Agca et al. 2015

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