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MicroRNAs (miRNAs) are small, highly conserved molecules that have been shown to regulate the expression of genes by binding to specific target mRNAs. Dicer, an RNase III endonuclease, is essential for the production and function of mature miRNAs and removal of Dicer has been shown to disrupt many developmental processes. In this report, Dicer was removed specifically from the retina using a floxed Dicer conditional allele and the retinal Chx10Cre transgene. Retinal Dicer knockout mice displayed a reproducible inability to respond to light. In addition, morphological defects were observed with the formation of photoreceptor rosettes at P16 which progressed to more general cellular disorganization and widespread degeneration of retinal cell types as the animals aged. This was accompanied by concomitant decrease in both scotopic and photopic ERG responses. Interestingly, removing a single allele of Dicer resulted in ERG deficits throughout life but not to morphological abnormalities. Northern blot analysis of Dicer depleted retinas showed a decrease in several microRNAs. The observation that progressive retinal degeneration occurred upon removal of Dicer raises the possibility that miRNAs are involved in retinal neurodegenerative disorders.
MicroRNAs (miRNAs) encode small 21-nt single-stranded RNAs that recently have been shown to regulate gene expression in a large number of diverse organisms and tissues (Cao et al., 2006). In vertebrates, miRNAs most often function as negative regulators of gene expression by base pairing with the 3’-untranslated region (3’-UTR) of target mRNAs while part of RNA-induced silencing complexes (RISCs). More than 400 miRNAs have been identified in both mice and humans, and each miRNA has the potential to regulate hundreds of target genes (Ambros and Cen, 2007; Lagos-Quintana et al., 2002). It has been proposed that more than one third of all human genes may be regulated by miRNAs (Lewis et al., 2005). Approximately 70% of experimentally detectable miRNAs are expressed in the brain, and among those upregulated in a tissue specific expression pattern in the embryo, half are brain-specific/enriched (Cao et al., 2006; Babak et al. 2004, Barad et al. 2004, Miska et al. 2004, Sempere et al. 2004, Thomson et al. 2004).
miRNAs are processed by Dicer, a double-stranded RNA-specific endonuclease, from a non-functional ~70-nt precursor RNA to a functional ~21-nt molecule (Ambros et al. 2003). Dicer null mice have been reported to die at embryonic day 7.5 (Bernstein et al. 2003) rendering it impossible to study the role Dicer may play in later stages of development or in adult tissues. To bypass the early lethality associated with the removal of Dicer in all tissues, we recently created a floxed conditional Dicer allele (Harfe et al., 2005). Using this allele in conjunction with tissue-specific cre alleles, we and others have inactivated Dicer in a number of discrete tissues during both embryonic and postnatal life (i.e. Harfe et al., 2005; Harris et al., 2006; O’Rourke et al., 2007). Since Dicer is required for the processing of miRNAs, removal of Dicer results in a significant decrease in functional miRNAs.
In spite of their abundance in the brain, detailed functional studies of miRNAs in the nervous system have not been provided until very recently (Cuellar et al., 2008; Davis et al., 2008). In this report we used the Dicer conditional mouse in combination with a retinal Chx10Cre transgenic line (Rowan and Cepko, 2004) to assess the role miRNA-mediated regulation of gene expression plays in the developing and adult murine retina.
Our data demonstrate that inactivation of Dicer in the retina results in progressive and widespread structural and functional abnormalities, culminating in loss of photoreceptor mediated responses to light and extensive retinal degeneration. This is the first study of miRNA inactivation within the mammalian eye and suggests that miRNAs play an essential role in neural tissue homeostasis. The degenerative phenotype reported here raises the possibility that miRNAs may be involved in some retinal diseases, and perhaps more generally, in neurodegenerative disorders as suggested by recent results obtained in mice with a conditional knockout of Dicer in Purkinje cells (Schaefer et al., 2007).
Construction and genotyping of the Dicer and Chx10Cre alleles has been described previously (Harfe et al., 2005; Rowan and Cepko, 2004). To create mice in which Dicer was inactivated in the retina, males containing the Chx10Cre transgene were crossed to females homozygous for the Dicer conditional allele. This cross generated animals in which one Dicer allele was removed in the retina (“heterozygous animals”). Heterozygous mice were then used to create Dicerflox/Dicerflox;Chx10Cre animals (referred to as “CKO”). CKO animals were viable and did not contain any obvious systemic phenotypic defects. Dicerflox/Dicerflox mice were used as wild type controls in all experiments. All animals used in our analysis were outcrossed at least three times onto a C57Bl/6 genetic background.
Dicerflox/Dicerflox;Chx10Cre animals were crossed to R26R mice (Soriano, 1999) to visualize cells that had undergone a recombination event.
Mice were dark adapted overnight in a Faraday shielded room and all subsequent procedures were carried out under dim red light (>650 nm). The corneas of the mice were given drops of 1% atropine, 2.5% phenylephrine and 0.5% proparacaine (Akorn, Buffalo Grove, IL) for mydriasis and topical corneal anesthesia. The mice were then anesthetized by intraperitoneal injection (6 µL per gram) of a sterile mixture of 100 mg/mL ketamine, 20 mg/mL xylazine, and normal saline at a 1:1:5 ratio, respectively. When the mice were fully sedated, they were placed onto a heated ERG platform in ventral recumbency. Balanced salt solution (Alcon, Ft. Worth, Tx) and Gonak (Akorn, Buffalo Grove, IL) were used to lubricate the cornea prior to electrode placement. Custom platinum loop electrodes were positioned on the cornea using a 3-axis micro-positioning system (Narishige, Japan). Grass platinum subdermal needle electrodes (Grass, West Warwick, RI) were utilized for ground and reference. All ERG’s were performed using a Multiliner Vision (Jaeger/Toennies, Hochberg, Germany) system equipped with a Ganzfeld-stimulator (Jaeger/Toennies, Hochberg, Germany). The Multiliner Vision apparatus complies with and exceeds the relevant International Society for Clinical Electrophysiology of Vision standards. Unless otherwise indicated, all mice were tested for dark-adapted ERG responses followed by light-adapted ERG responses. White light from a xenon source was used as the stimulus. For the scotopic (dark-adapted) ERG series, mice were exposed to a series of 10 flashes at 0.1 cd•s/m2. For the photopic (light-adapted) ERG series, mice were exposed to background light at 100 cd/m2 for 1 minute and ERG recordings were done in the presence of a constant 100 cd/m2 background light. Mice were then exposed to a series of 50 flashes at 10 cd•s/m2. Right and left eyes were tested simultaneously and the data recorded. Analysis of the ERG waveforms for a-and b-wave maxima was carried out using the Multiliner Vision software. Further analysis was performed using Sigma Plot software. When calculating significance a Student’s t-test was used.
Retinas from 1 month (n=3, mutant; n=6, wt), 3 months (n=3, mutant; n=2 wt) and 24 months (n=2 mutant; n=3 heterozygous; n=2 wt) animals were pooled. RNA was isolated from fresh homogenized pooled retinas with TRI Reagent (Sigma). Approximately 12 µg of total RNA for the 1 and 3 month time points and ~8 µg of total RNA from 24 month animals was resolved on 10% urea/polyacrylamide gels and electroblotted to Hybond N+ membranes (Amersham) at 200 mA for 3 h. Blots were crosslinked using a Stratalinker (Stratagen) and prehybridized for at least 1 h at 37°C in ULTRAhybTM-Oligo (Ambion) hybridization buffer before overnight incubation at 37°C in hybridization buffer containing [32P]-end-labeled probe. Probes were generated by end-labeling 20 pmols of DNA oligonucleotide (Invitrogen) complementary to a specific miRNA or U6 with T4 polynucleotide kinase (New England Biolabs) and 250 µCi [γ-32P] ATP (Perkin Elmer) followed by purification with MicroSpinTM G-25 columns (Amersham). Blots were washed (2X SSC, 0.1% SDS) at 37°C for 30 min followed by two 30 min room temperature washes. Blots were exposed to Kodak BioMax film and quantification was determined by densitometry of autoradiographs using ImageQuant TL v2003.02 software. For subsequent probing, blots were stripped by incubating twice in boiling 1% SDS for 15 minutes each and exposed to film to confirm that probes were removed.
Whole mouse eyes were fixed overnight in 4% PFA/PBS, infiltrated with 30% sucrose, frozen and sectioned as described below. A 260-nt riboprobe (Harris et al., 2006), specific to the floxed Dicer exon was used to assess Dicer distribution. After a brief wash in PBS, retinal sections were treated 2×10 min with RIPA buffer, post-fixed 15 min in 4% PFA /PBS and acetylated with 0,25% acetic anhydride in triethanolamine 0,1 M. Slides were then pre-hybridized for 2 hrs in a solution of 50% formamide, 5× SSC, 5× Denhart’s, 500 µg/ml salmon sperm DNA and 250 µg/ml yeast RNA. Slides were hybridized for 12 hrs at 48 °C, with 100–300 ng of digoxigenin labeled probe per slide in a plastic chamber humidified with 50% formamide in 5× SSC. Post-hybridization washes were made with 50% formamide, 0,1% Tween-20 in 2× SSC, for 2 hrs. Sections was then re-equilibrated in 0.1 % Tween-20 in 0.1M maleic acid buffer (MABT), blocked for 1–2 hours in 10 % sheep serum in MABT and incubated for 12 hrs at 4°C with anti-digoxigenin-AP (1/2000 in blocking buffer; Roche). After MABT washing, the pH of the specimens was adjusted with 100 mM Tris pH 9,5, 50 mM MgCl2, 100 mM NaCl. Finally, retinal sections were reacted for 1–2 hrs with NBT-BCIP solution (Sigma), washed in PBS, dehydrated in ethanol, mounted in DPX and acquired with an Zeiss Axioplan microscope, equipped with an AxioCam HRC color camera and dedicated AxioVision software.
Three intraperitoneal (IP) injections of 50 mg/Kg BrdU were performed at 3 hr intervals in P35 mice (n=5). Mice were then sacrificed 3 hrs after the final injection, their eyes prepared as described above and BrdU detected by immunocytochemistry on frozen retinal sections following the procedure of Close et al., 2005.
The eyes of mice aged postnatal (P) 2, 16, 30, 45 days and 3, 4, 5, and 7 months were harvested, quickly enucleated and immersion-fixed for 1 hr in 4% paraformaldehyde in 0.1 M phosphate buffer (0.1M PB). The eyes were then rinsed in buffer, infiltrated overnight in 30% sucrose in 0.1M PB, embedded in OCT/Tissue Tek (Sakura) and frozen on a cryostat stage at −25/−30°C. Eyes were sectioned vertically in 12–16 µm serial sections with a Leica cryostat. Sections were collected on Superfrost Plus slides and air dried for 5 min to 2 hours. Slides were rinsed for 10 min with 0.01M PBS and blocked for 2 hours in a solution containing 5% bovine serum albumin (BSA) and 0.3% Triton-X100 in PBS. Primary antibodies were diluted in 1% BSA, 0.1% Triton X100 in PBS. Slides were incubated in primary antibodies for 12–18 hours at 4°C. Primary antibodies were rinsed off by 2×10 min incubation in PBS at RT. Sections were then incubated for 2–4 hrs in solutions containing appropriate secondary antibodies, diluted 1:400, conjugated with Oregon Green 488, Alexa Fluor 568, Alexa Fluor 647 (Invitrogen) or Cy-3 (Sigma). Sections were then rinsed in PBS and, if appropriate, counterstained with the fluorescent nuclear dyes BOBO-1 or TOTO-3 (Invitrogen).
Primary antibodies and dilutions were as follows: mouse anti-rhodopsin (1:2,500; Sigma); rabbit anti-recoverin (1:2,000; Chemicon); mouse and rabbit anti-protein kinase C-α (PKC-α; 1:1,000; clone MC5, Sigma; sc-208, Santa Cruz Biotechnology); mouse and rabbit anti-calbindin D-28k (1:2,000; clone CB955, Sigma; Swant); mouse anti-G0α (1:1,000; MAB 3073, Chemicon); mouse anti-neurofilament 200 kDa (1:100; NF-200; clone N52, Sigma); rabbit anti-mGluR6 (1:2,000; from Dr. S. Nakanishi, Osaka University, Japan); mouse anti-post-synaptic density protein 95 (1:500; PSD95; AbCam); mouse anti-glutamine synthase (GS; 1:2,000; MAB302, Chemicon); rabbit anti-glial fibrillary acidic protein (1:1,000; GFAP, Sigma); goat anti-choline acetyl transferase (1:500; ChAT; Chemicon); mouse anti-Cre recombinase (1:1,000; MAB3120; Chemicon); rabbit anti-Green Fluorescent Protein-Alexa Fluor 488 conjugate (1:1,000; Invitrogen); rabbit anti-phosphohistone H3 (1:400; Upstate); rabbit anti atypical PKC (C20, 1:500, Santa Cruz Biotechnology); rabbit anti zonula occludens-1 (ZO-1; 1: 200, Zymed); rabbit anti laminin (1:400, Sigma); guinea pig anti vesicular glutamate transporter (vGLUT1, 1:500, Chemicon); rabbit anti vesicular GABA transporter (vGAT 1:500, Synaptic System). Rabbit anti S-cone opsin (1:1000, Chemicon) and Alexa Fluor 488-PNA lectins (1:400; Invitrogen) were used to label cone photoreceptors.
Retinal preparations were examined with a Leica TCS-NT confocal microscope equipped with an argon-krypton laser or with a Leica TCS-SL spectral confocal microscope using high numerical aperture oil immersion objectives. Images acquired at a resolution of 1024×1024 were saved as TIFF files and exported on a workstation for offline analysis. Retinas from at least one wild type, one heterozygote and three CKO littermates for each age group (2, 16, 30, 45 days and 3, 4, 5 and 7 months) were used for ICCH and screened with the full panel of antibodies listed above. A total of 57 animals were analyzed for ICCH. Except for P2 and P16 animals, eyes used for morphological analysis were from animals previously used for ERG recordings (see below).
One additional litter composed of 4 mutants and 5 wild type animals aged P2 was used for EM studies of immature retinas. After decapitation, the eyes were removed and immersion-fixed in 2% PAF and 2.5% glutaraldehyde for 12 hrs. After dissections, retinal tissue was postfixed in osmium tetroxide, bloc stained with 1% uranyl acetate, dehydrated in ethanol and embedded in plastic. Semithin sections (1–2 micrometer thick) were stained with Epoxy Tissue stain (EMS) and observed at the light microscope. Ultrathin sections from retinal blocs were counterstained with uranyl acetate and lead citrate and examined with a Jeol 1200 EXII electron microscope. Photographs of retinal neuroblasts and differentiating photoreceptors were taken at 8,000–20,000X.
An analysis of electroretinograms (ERG) recorded from 1, 3 and 5 month old CKO, heterozygous and wild type eyes revealed that the scotopic a- and b-wave (Fig.1A) and photopic b-wave (Fig.1B) amplitudes were diminished in both the CKO and heterozygous animals, with the CKO most severely affected. CKO eyes had the lowest amplitudes in all ERG tests and were significantly different from wild type (Fig.2A,B and Table 1). When compared to heterozygous eyes, all but two ERG measurements were significantly different (Fig.2A, B and Table 1). As a group the heterozygous eyes were significantly different from WT in all but one measurement (Fig.2A, B and Table 1). Both CKO and heterozygous eyes showed variability in the level of attenuation, consistent with the mosaic expression of the Chx10Cre allele (see below). The observed decrease in ERG function in heterozygous mice was unexpected, based on the lack of any detectable immunohistochemical defect in these animals. These data suggest that the proximal cause of such a functional abnormalities may reside in molecular changes affecting phototransduction rather than retinal morphology.
Given the profound abnormality of ERG responses in all the CKO animals tested, and their progressive inability to respond to light, we carried out a morphological analysis of their retinas attempting to establish a correlate between abnormal function and anatomical phenotype.
As previously reported (Rowan and Cepko, 2004), Cre expression from the Chx10Cre transgenic allele does not drive cre expression in all retinal cells. As a result, not all cells in CKO retinas would be expected to lack a functional Dicer protein. To determine which retinal cells in CKO animals lacked Dicer, we performed RNA in situ hybridizations on retinal sections using a probe that was specific for the floxed exon in the Dicer conditional allele (Harris et al., 2006). In wild type retinas, native Dicer was expressed in the vast majority of retinal cells (Fig.3 A). Staining in photoreceptors appeared localized in the inner segments. Some cells of the inner nuclear layer and ganglion cell layer were also labeled. Ganglion cells, possibly because of their abundant cytoplasm, appeared more intensely stained than other cell types.
In the P16 CKO retinas (Figure 3B), Dicer expression was found to be patchy, with groups of labeled cells interdigitated with unlabeled cells. For example, cells in the ganglion cell layer were negative in some areas and positive in adjacent regions (Fig.3B). To determine if patchy expression of Dicer in CKO retinas was a reflection of the mosaic expression of the Chx10Cre driver, we examined Cre expression in CKO animals by means of anti-Cre antibodies. In the retinas of these mice, nuclei of cells in the inner nuclear layer, most likely bipolar cells, were stained as expected (Figure 4). However, the pattern of labeling was not uniform but contained patches of Cre-negative cells contiguous to clusters of Cre-positive cells.
The cre gene is fused to GFP, and the fusion is run off the chx10 promoter (Rowan and Cepko, 2004). Therefore, we could also reveal cre-positive cells using anti-GFP antibodies. The pattern of staining obtained with either Cre or GFP antibodies was identical (data not shown). The mosaic expression of Dicer in the CKO retinas was also shown by staining for β-Gal on retinal sections obtained from mice containing the R26R cre inducible reporter allele (Chx10Cre; R26R; Dicerflox/Dicerflox). The R26R allele allowed for the detection of cells that had undergone a recombination event due to expression of cre recombinase from the Chx10Cre transgene (Soriano, 1999) (see supplemental figure S1).
Our data indicate that Dicer inactivation using the Chx10Cre transgene results in the production of mosaic retinas in which Dicer null cells are located adjacent to Dicer positive cells.
To determine if partial Dicer inactivation in CKO retinas resulted in aberrant miRNA processing, we performed a small RNA northern blot for miRNAs known to be expressed specifically in the retina or elsewhere in the CNS (miR-96, miR-124a and miR-204). miR-96 and miR-204 are highly expressed in the developing and adult retina, while miR-124a is expressed throughout the entire CNS (Deo et al., 2006; Karali et al., 2007; Ryan et al., 2006). Northern blots were performed on pooled retinas from control and CKO animals (see Methods).
At 1 month, compared to wild type, the mature form of all three miRNAs examined was not changed (Fig. 5). At three months, mature miR-96 and miR-124 were found to be decreased 70% compared to wild type. At two years, mature miR-96 was decreased 63% and miR-124 was decreased 36%. Interestingly, we observed a decrease in production of mature miR-96 and miR-124 in heterozygous retinas (Fig. 5). The decrease in miRNA levels in heterozygous retinas does not result in a phenotypic defect (see Fig.10B) but may, at least partially, explain why heterozygous animals contained defective ERGs (see Fig. 1 and Fig. 2). A decrease in the number of retina cells, compared to wild type, expressing the retinal-specific microRNA miR-183 was also detected in 7 month-old CKO retinas by means of RNA in situ hybridization (see supplemental figure S2).
At all stages examined, miR-204 expression was unchanged (Fig. 5). miR-204 has been reported to be specifically expressed in a subset of cells in the inner nuclear layer of the retina (Deo et al., 2006). The Chx10cre allele used in our experiments is expressed in a very mosaic pattern in the inner nuclear layer (see Fig. 4), which may be the cause of our inability to detected alterations in the expression of this particular miRNA in mutant animals. Both miR-124 and miR-96 have been reported to be broadly expressed throughout the retina (Karali et al., 2007; Xu et al., 2007). Upon removal of Dicer from the retina, cells undergo progressive cell death (see Fig. 9D and Fig. 10A). The increase in mature miRNAs found in Dicer null retinas at the two year time point compared to three months could be due to a decrease in the number of Dicer null cells present in the retina at this late stage.
The most striking abnormal feature present in all CKO retinas aged P16 and older was the presence of photoreceptor rosettes (Figure 6). Rosettes are circular structures, comprising photoreceptors only, that are oriented toward an internal lumen with photoreceptor outer segments protruding inward. Fragments of retinal pigment epithelial cells are also occasionally observed inside rosette lumen (not shown).
In P16 mutants, rosettes were scattered along an otherwise normal retinal outer surface and were primarily composed of photoreceptors and their synaptic terminals (Fig. 6). The number and density of rosettes varied in each retina, however each rosette was separated from the others and clearly identifiable even at low magnification. Interestingly, the presence of rosettes did not appear to alter the typical laminar organization of the retina: inner retinal layers were unperturbed by overlying rosettes and the number of rows of photoreceptor nuclei in areas devoid of rosettes was found to be normal (12–14; data not shown). At this early stage, the presence of rosettes was not accompanied by concomitant overexpression of glial fibrillary acidic protein (GFAP) in Müller cells. GFAP overexpression is a well known indicator of glial activation and is associated with retinal degeneration or perturbation in laminar organization (Marc et al., 2003). In the CKO animals, GFAP overexpression in Müller cells was only detected at later stages during which a clear loss of retinal cells was taking place (Fig 9E).
Rosettes have previously been described in a number of pathological retinal conditions, including retinoblastoma (Yuge et al., 1995); diabetic retinopathy (Lahav et al., 1975) and retinitis pigmentosa (Tulvatana et al., 1999). In these diseases, retinal degeneration and/or abnormal proliferation are present (Lin et al., 2001). To test whether the formation of photoreceptor rosettes in Dicer depleted retinas was associated with an increase in cell proliferation in the photoreceptor layer we treated P35 animals with BrdU. In addition, we stained sections from multiple P16 retinal samples with antibodies against phosphohistone H3, a specific marker of cell proliferation (Dhomen et al., 2006; Fig. 7). Using either method, we failed to detect dividing cells in CKO retinas, although we observed, as expected, mitotic cells in other ocular tissues such as the cornea (Fig. 7). Thus rosette formation did not appear to be caused by an upregulation in retinal cell proliferation.
To determine if areas of rosette formation were those that had experienced Cre expression, tissue samples were labeled with anti-Cre antibodies. Rosettes were found in areas where bipolar cells expressed cre suggesting that these structured formed as a result of a decrease in miRNA levels (Fig.7).
Appropriate formation of retinal layers depends critically upon the maintenance of apico-basal polarity in the neuroepithelial cells during early development. Retinal neuroblasts are joined by junctional complexes precisely positioned at the apico-basolateral domain of the cell membrane. Alterations of the molecular components controlling these complexes lead to abnormal retinal layering and subsequent rosette formation (Erdmann et al., 2003; Malicki et al., 2003; Masai et al., 2003; reviewed in Malicki, 2004; Galli-Resta et al., 2008). To test the hypothesis that the rosettes observed in Dicer CKO mature retinas could be caused by major defects in the apico-basal polarity of neuroepithelial cells, we examined by ICCH and electron microscopy (EM) CKO and wt retinas at 2 days of age (n=8 animals for each genotype). We assessed a) early retinal layering; b) laminar position of dividing and differentiating cells and; c) position of junctional complexes among differentiating photoreceptors and neuroblasts. Results are illustrated in supplemental figure S3 and supplemental figure S4.
This analysis demonstrated that immature Dicer CKO retinas did not contain major defects in neuroepithelial cell polarity or in the structure of adherens junctions, therefore indicating that rosettes, which are found in older mutant retinas, were not the result of improper patterning of these cell types.
The normal complement of retinal cell types, comprising neurons and glia, (i.e. Jeon et al., 1998; Haverkamp and Wassle, 2000, etc.), were represented in the CKO retinas from P16 onwards. Cells were observed to occupy the expected laminar positions and the overall retinal architecture appeared normal. One exception, as discussed above, was the presence of rosettes at all stages examined. Using cell-type specific antibodies all major cell types including rods and cones (Fig.8A), horizontal cells (Fig.8B), rod and cone bipolar cells (Fig.8C, D), various types of amacrine cells (Fig.8E) and ganglion cells (Fig.8F) were present in CKO retinas. All cells displayed normal morphology, pattern of stratification and appropriate lamination in the plexiform layers. Retinal glial cells, in particular astrocytes and Müller cells (Fig.7A and B), were also appropriately represented and positioned correctly in the CKO retinas.
The majority of rod bipolar cells were Cre-positive, as demonstrated by anti-Cre (Fig.8C) and anti-GFP antibodies (not shown). As expected, Cre staining was nuclear and showed a similar pattern in the CKO and wt retinas (Fig.4A and B). Cre-positive nuclei occupied the presumptive bipolar layer, and thus both rod and cone bipolar cells were Cre-positive and presumably Dicer null. A few nuclei from Cre-positive, PKCα-negative cells were larger in size and were presumably derived from Müller cells.
Using cell type specific antibodies, the data suggested that removal of Dicer in retinal progenitors and differentiated bipolar cells did not affect retinal cell fate or migration of cells to their wild type locations during retinal development.
The time period from P16 to P45 was characterized by progressive alteration and remodeling of the laminar retinal structure in CKO mice (Fig. 9). The number of rosettes increased (Fig.9A) while their structure became more complex: bipolar and horizontal cells penetrated the rosettes while photoreceptor clusters were displaced toward the outer retina (Fig. 9). The normal synaptic structures of the OPL also became displaced toward the outer retina (Fig.9A, B); concomitantly, the dendrites of both rod bipolar and horizontal cells sprouted profusely in the direction of the photoreceptors (Fig.9C–E). Entire retinal domains appeared to be displaced outward, resulting in a progressive destruction of the external lamination of the P45 CKO retina. Most dramatically, PSD95, a marker of photoreceptor synaptic terminals, was expressed ectopically in the outer retina and reached the outer limiting membrane (Fig.9A).
As CKO animals reached 3 months of age, the rosettes were observed to decrease in size and eventually disappear from the retina. Photoreceptors composing these rosettes degenerated and the outer nuclear layer became progressively thinner. This was confirmed by detection of pycnotic nuclei in the outer and inner retina by using DNA-binding dyes (Fig.10A). At 3 months, most rosettes had been lost and replaced by areas of extensive degeneration. Generally, the outer nuclear layer appeared to be more reproducibly thinner in the peripheral as compared to the central retina, suggesting that the degenerative processes had a periphery-to-central gradient. At 3 months of age, in the extreme periphery, the outer nuclear layer was reduced at 2–3 rows (Fig.9F).
Concomitant with outer retinal degeneration, the inner retina also deteriorated and exhibited cellular loss. Simultaneously, Müller cells became hypertrophic with their radial processes increasing in size and GFAP reactivity raising visibly, indicating a generalized glial activation (Fig.9F). At the same time, Cre positive cells became scarce, most likely because a large fraction of these Dicer null cells degenerated, as demonstrated by the progressive thinning of the inner nuclear layer. Surviving rod bipolars often occurred in clusters and had altered morphologies with axonal arborizations that were hypertrophic with enlarged, bulbous like endings and dendrites that were scant and thicker than normal. PKCα staining was abnormally bright and had an unusually punctate appearance. Most of the residual rod bipolar cells were Cre-negative (Fig.9D) and therefore most likely originated from precursors in which Dicer was not inactivated due to the mosaic activity of the Chx10Cre transgenic allele (Rowan and Cepko 2004).
Retinas of animals in which a single copy of Dicer was inactivated in the retina (heterozygous animals) were examined by in situ hybridization, immunocytochemistry and detailed confocal microscopy. Using these assays, no major defects were detected (Figure 10). However, ERG recordings and Northern blot analysis (see above) did reveal functional abnormalities and a decrease in retinal miRNAs.
The role miRNAs play in the development and maintenance of the nervous system is unknown. Dicer-deficient zebrafish exhibit abnormal morphogenesis and widespread abnormalities in neural development (Giraldez et al. 2005). However, many aspects of early embryonic development, including patterning and cell fate specification are largely unaffected by Dicer deletion in zebrafish, in part due to the inheritance of maternal Dicer. Dicer-deficient mice die during early development, around embryonic day 7.5, and were found to be depleted of pluripotent embryonic stem cells (Bernstein et al. 2003). In the mouse nervous system, depletion of Dicer in Purkinje cells has been reported to result in neuronal degeneration (Schaefer et al., 2007).
The results reported here highlight the crucial role Dicer and miRNAs play in the long-term regulation of retinal cell lamination, survival and function. We have shown that removal of Dicer in the mouse retina had no visible impact on early postnatal retinal structure and function since retinal lamination appeared normal and all expected retinal cell types were represented. In addition, the relative widths of the various retinal layers appeared to be very similar between CKO and wild type retinas, although the perturbation to outer retinal surface in the vicinity of photoreceptor rosettes and the mosaic expression of the Dicer allele with alternation of normal and abnormal retinal patches, made it virtually impossible to perform histograms of cell distributions. However, by 1 month of age the response to light was significantly diminished and an abnormal retinal lamination clearly detectable.
It is important to note that the Cre allele used in these experiments, Chx10Cre, drives Cre expression in the retina during embryogenesis. In both lung (Harris et al., 2006) and limb (Harfe et al., 2005) loss of Dicer resulted in a substantial decrease of all miRNAs examined ~2 days after Cre expression. In our experiments in the retina, Dicer was removed more than 20 days prior to ERG analysis and morphological detection of rosettes at P16.The presence of mature miRNAs at wild type levels 1 month after birth indicates that either miRNAs in the retina are extremely stable or that an additional protein can compensate for DICER function in this tissue during early postnatal life. This is in stark contrast to what was observed in the mouse limb where Dicer removal almost immediately resulted in massive cell death (Harfe et al., 2005). The persistence of mature miRNAs in the Dicer CKO retina might explain the late degeneration of retina cells observed in our experiments.
As CKO animals aged, we observed numerous defects in retinal structure including cell death. These observations suggest that miRNAs are initially required for the retina’s ability to respond to light, but not for cell survival. However, miRNAs play an essential role in maintaining the majority of cell types found in the adult retina.
Surprisingly, the light response deficit was observed in heterozygous in addition to homozygous animals suggesting that the level of Dicer protein was essential for proper function of the mouse retina. The light response defect was observed in all ages of retinas lacking a single Dicer allele although we could detect no morphological defects in the retina of these heterozygous animals at any stage of life. These data suggest that either the mouse retina is extremely sensitive to the amount of mature miRNAs present or that Dicer has additional, non-miRNA functions in the mouse retina (see below). To our knowledge, this is the first demonstration in which the dosage of Dicer protein within a cell influences function. This light response deficit seen in heterozygote animals suggests one or more elements of the phototransduction cascade itself may be affected, since alteration here would likely quickly affect the ERG response but not retinal structure in the short term.
It is possible that not all the phenotypes we observed upon removal of Dicer in the retina were due to the loss of mature miRNAs. Dicer could be processing other double-stranded RNAs besides miRNAs within the targeted retinal cells. In addition, this enzyme could play a role in the nucleus as suggested by loss of centromeric and pericentromeric silencing in Dicer null ES cells (Murchison et al., 2005; Kanellopoulou et al., 2005).
Given the fact that the first sign of morphological abnormality in CKO retinas was the formation of photoreceptor rosettes and that photoreceptor function was affected early, our data suggest that photoreceptors are the retinal cell type most sensitive to an imbalance in the production and maturation of miRNAs.
Photoreceptor rosettes are a common finding in a variety of retinal diseases. In particular, they are found in diabetic retinopathy, retinitis pigmentosa and ischemia (Lahav et al., 1975; Yuge et al., 1995; Tulvatana et al., 1999). Rosettes have also been reported to result from decreased levels of secreted proteins of the Wnt family, shown to function as organizers of retinal lamination. Rosette formation is usually associated with alteration of cytoskeleton, abnormal production of extracellular matrix and adhesion molecules (Li and Sakaguchi, 2004; Lunardi et al., 2006) in a variety of vertebrates, including Xenopus, zebrafish, chicken, mice and humans.
Despite the fact that the exact trigger of rosette formation is not known, this common abnormality in retinal lamination might be caused early in development through alterations in polarity of neuroblasts (reviewed in Malicki, 2004). To test this hypothesis, we compared wt and Dicer CKO retinas at two days of age, a time at which inner retinal layers are partially formed and the outer retina is still largely composed of neuroblasts and differentiating photoreceptors (Sharma et al., 2003). Morphological examinations of retinal sections by ICCH and EM, aimed at assessing the proper orientation and polarity of neuroblasts, mitotic cells, developing photoreceptors and laminar position of differentiated cells, did not reveal major abnormalities. Adherens junctions in particular were found at their appropriate location at the outer retinal margin between differentiating photoreceptors and neuroblasts. These data were consistent with our observation that wild type amounts of miRNAs were found during early postnatal development. Hence, it appears that rosette formation in the Dicer KO retina is not linked to major changes in neuroblast polarity or cell orientations and instead occurs upon completion of retinal development.
Rosette formation may be followed by progressive photoreceptor death (Lin et al., 2001). This is similar to what we observed upon removal of Dicer in the retina. While the pattern of cell degeneration observed in mouse Purkinje cells upon Dicer inactivation (Schaefer et al., 2007) bears similarity to neurodegenerative disorders such as Alzheimer’s and Parkinson’s disease, the pattern of retinal degeneration reported here does not precisely parallel genetically well understood forms of inherited retinal disease, such as Retinitis Pigmentosa. In both the human disease and in mouse models of this disorder, a clear rod-cone degeneration is observed first, while the degeneration of inner neurons, including horizontal and bipolar cells, is usually secondary (Gargini et al., 2007). This partially reflects the fact that the primary genetic defects typically occurs in a photoreceptor-specific gene. In the Dicer CKO retina examined here, cellular death after P30 affected both rod and cone cell function simultaneously and was accompanied by a concomitant decrease in the number of second order neurons. Interestingly, rod bipolar cells that persisted in retinas aged 3 months and older were also Cre-negative. Albeit their morphology was abnormal, the fact that these cells survived longer than Dicer null rod bipolars suggests the occurrence of cell autonomous events in the genesis of the complex CKO phenotype.
At least 78 of the >400 miRNAs reported in mice are expressed in the retina (Xu et al., 2007). Since each miRNA could potentially regulate hundreds of genes, it is unclear what the role individual miRNAs may play in retinal development during embryonic and postnatal life. To unravel the exact role miRNAs play in rosette formation and/or ERG loss, the conditional removal of miRNAs specifically expressed in the retina, such as miR-183 (Karali et al., 2007), could be attempted.
Inactivation of Dicer in the mouse model used here likely resulted in a decrease in global levels of most retinal-expressed miRNAs. The unique pattern of retinal degeneration uncovered in this report is the first step towards identifying the overall role miRNAs contribute to retinal cell function and survival. Our results provide insight into the molecular mechanisms of retinal degeneration and the maintenance of normal retinal function and architecture.
Funded by grant NIH R01 EY12654 (ES); EY011123, EY008571, EY13729, EY007132 and NS36302 and grants from the MVRF, FFB and JDRF, and from RPB (WH & JA); Howard Hughes Medical Institute and NIH EY0 9676 (CC); NIH Grant R03-DA22201(MTM); UF start-up funds (BDH).