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. Author manuscript; available in PMC: 2017 Sep 1.
Published in final edited form as: Exp Hematol. 2016 Jun 1;44(9):826–837.e4. doi: 10.1016/j.exphem.2016.05.012

The Spliceosomal Component Sf3b1 is Essential for Hematopoietic Differentiation in Zebrafish

Adriana De La Garza 1,2,#, Rosannah C Cameron 1,2,#, Sara Nik 1,2, Sara G Payne 1,2, Teresa V Bowman 1,2,3,#
PMCID: PMC4992596  NIHMSID: NIHMS792063  PMID: 27260753

Abstract

SF3B1 (Splicing factor 3b, subunit 1) is one of the most commonly mutated factors in myelodysplastic syndrome (MDS). Although the genetic correlation between SF3B1 mutations and MDS etiology are quite strong, no in vivo model currently exists to explore how SF3B1 loss alters blood cell development. Using zebrafish mutants, we show that proper function of Sf3b1 is required for all hematopoietic lineages. Similar to MDS patients, zebrafish sf3b1 mutants develop a macrocytic anemia-like phenotype due to a block in maturation at a late progenitor stage. The mutant embryos also develop neutropenia as their primitive myeloid cells fail to mature and turn on differentiation markers such as l-plastin and myeloperoxidase. In contrast, production of definitive hematopoietic stem and progenitor cells (HSPCs) from hemogenic endothelial cells within the dorsal aorta is greatly diminished, while arterial endothelial cells are correctly fated. Notch signaling, imperative for the endothelial-to-hematopoietic transition, is also normal, indicating HSPC induction is blocked in sf3b1 mutants downstream or independent of Notch signaling. The data demonstrate Sf3b1 function is necessary during key differentiation fate decisions in multiple blood cell types. Zebrafish sf3b1 mutants offer a novel animal model to explore the role of splicing in hematopoietic development and provide an excellent in vivo system to delve into the why and how Sf3b1 dysfunction is detrimental to hematopoietic differentiation, which could enlighten MDS diagnosis and treatment.

Introduction

Transcription and RNA processing are interconnected processes that are essential for regulating gene expression. Genes involved in these processes are commonly mutated in hematological malignancies. Although many key transcription factors involved in both embryonic and adult hematopoiesis have been identified1, the role of RNA processing factors is unclear.

Hematopoietic specification occurs in multiple waves. The first or primitive wave of hematopoietic induction consists mainly of erythrocytes and myeloid cells that sustain the embryo until adult cells are formed2. The definitive wave, which occurs later in development, generates multipotent progenitors and hematopoietic stem and progenitor cells (HSPCs) that give rise to all the mature blood cells required throughout the lifetime of an organism3. The first HSPCs emerge from hemogenic endothelium in the ventral wall of the dorsal aorta and then colonize secondary organs, such as fetal liver, placenta, thymus and bone marrow in mammals and the caudal hematopoietic tissue, thymus, and kidney marrow in zebrafish (reviewed in1-3). Much is known about the transcription factors, such as runx1 (runt-related transcription factor 1), gata2, scl (stem cell leukemia), gfi1 (growth factor inducible-1), pu.1, and gata1 among others, that orchestrate developmental hematopoietic specification, but less is known about the role of RNA processing.

Splicing of pre-mRNA is a co-transcriptional event that diversifies the proteome and contributes to cell fate control. The spliceosome is a macromolecular machine comprised of five snRNPs (small nuclear ribonucleoproteins), each made of large, multicomponent protein complexes and snRNAs (small nuclear RNAs). Recent genome sequencing efforts identified mutations in several spliceosomal components in hematologic malignancies, such as acute myeloid leukemia and myelodysplastic syndrome (MDS)4-8. These clinical data suggest splicing regulation is important in hematopoiesis. Mutations in spliceosomal components are mutually exclusive in MDS, yet mutations in each factor correlate with a different disease phenotype. For example, mutations in the U2 snRNP component SF3B1 (Splicing factor 3b, subunit 1) show a strong correlation with subtypes of MDS with refractory anemia and ringed sideroblasts9. In contrast, mutations in the splicing accessory protein SRSF2 (Serine/arginine-rich splicing factor 2) or U2AF1 (U2 snRNP-associated factor-1) correlate with poor prognosis and are found more frequently in more malignant forms of MDS, such as refractory anemia with excess blasts4-8,10. These clinical data imply that splicing factors might employ different mechanisms and/or have different cell-type selective tropisms that lead to distinct MDS features. Thus, it is imperative to understand the specific roles of each splicing subunit.

In support of these clinical observations, zebrafish spliceosomal mutants display some overlapping and distinct defects in hematopoiesis11-16. For example, mutants in the RNA helicases ddx18hi1727 (DEAD-box protein 18) and dhx8hg9 (DEAH-box protein 8) have greatly reduced primitive myeloid and erythroid cells, while ddx46hi2137 (DEAD-box protein 46) mutants have impaired primitive erythropoiesis as well as defects in definitive HSPCs maintenance11-13. In contrast, mutants in the U5 snRNP component prpf8gl1 (pre-mRNA processing factor 8), have defective myeloid lineage development, but no defects in erythropoiesis or definitive HSPC induction14. The U2 snRNP-associated factor U2af1 is also critical for hematopoietic development, as zebrafish u2af1hi199 mutants have diminished erythroid differentiation and fewer HSPCs16. In contrast, mutants for the U1 snRNP component snrnp70hi3018 (small nuclear ribonucleoprotein, 70kDa) and the U2 snRNP member sf3a3hi1950 (splicing factor 3A, subunit 3) show diminished HSPC formation, but no defects in primitive hematopoiesis15. These data show that dysfunction in numerous components of the spliceosome negatively impact hematopoiesis, but also demonstrates differential cell-type sensitivity depending on the spliceosomal constituent mutated.

SF3B1 is the most commonly mutated spliceosomal component in MDS4-10, yet its in vivo consequences on developmental hematopoiesis are unknown. Homozygous Sf3b1-null mice are embryonic lethal pre-implantation, precluding analysis of development17. Adult heterozygous Sf3b1 mutants develop macrocytic anemia and have HSC self-renewal defects, demonstrating cell-type selective susceptibilities to Sf3b1 dosage18-20. To explore the in vivo outcomes of sf3b1 depletion on developmental hematopoiesis, we characterized a recently described zebrafish sf3b1 mutant21,22. These mutants contain a loss-of-function mutation that is partially compensated by retention of maternally-deposited wild-type protein and mRNA21. The sf3b1 mutants show increased neuronal cell death, similar to many other spliceosomal mutants, and have defects in neural crest cell induction21. Using the sf3b1hi3394a loss-of-function allele, we defined an essential role for sf3b1 in blood development. The primitive erythroid and myeloid cells are specified normally, but then fail to mature. In the definitive wave, HSPCs do not form, although the precursor endothelial cells are present. Notch signaling, a critical mediator of hemogenic endothelium and HSPC induction, also appears unaffected in the dorsal aorta in sf3b1 mutants. These data indicate sf3b1 is required during hemogenic specification downstream or independent of Notch. Overall, our findings show sf3b1 function is critical for key hematopoietic cell fate decisions during early development and establish sf3b1 zebrafish mutants as an in vivo model to delineate how Sf3b1 dysfunction alters hematopoietic decisions, which could impact MDS diagnosis and treatment.

Materials and methods

Zebrafish

Zebrafish were maintained as described.23,24 All fish were maintained according to IACUC approved protocols in accordance with Albert Einstein College of Medicine research guidelines.

Zebrafish lines used

The sf3b1 mutant line hi3394a contains a viral insertion between the 1st and 2nd exons of the gene sf3b1, resulting in a premature stop codon in exon 221,22. Wild-type and heterozygous embryos are phenotypically indistinguishable from one another, thus for all of the experiments presented in the paper, wild-type and sf3b1 heterozygous embryos are cumulatively referred to as wild type. As sf3b1 homozygous mutants are developmentally delayed and show defects in melanocyte development21, mutant embryos were age-matched for all analyses according to morphological features. In addition, analyses of each hematopoietic population were done for at least two developmental time points to control for any variation in stage matching. Mutant embryos die between 2-3dpf, precluding analysis of later stages21. For fluorescent imaging and flow cytometry analysis, we crossed sf3b1hi3394a heterozygous animals to the following transgenic lines: Tg(gata1:gfp)25, Tg(kdrl:dsRed)26, Tg(cmyb:gfp)27, Tg(lyz:dsRed)nz50 28 and Tg(tp1bglob:eGFP) (tp1:eGFP)29.

Morpholino injections

To knockdown sf3b1 levels, we utilized a published translation-inhibiting morpholino21. Wild-type embryos were injected with 0.1ng (0.5 nl of 0.2μg/μl stock) and compared to non-injected sibling controls.

Whole-mount in situ hybridization

In situ hybridization steps were performed as described previously by Thisse et al.30 with minor modifications: before proteinase K permeabilization, embryos older than 28 hours post fertilization (hpf) were bleached after re-hydration to remove pigmentation. The bleaching was done for 5-10 minutes using a bleaching solution of 0.8% KOH, 0.9% H2O2 and 0.1% Tween 20. The following probes were used: β-globin31, ephrinb232, flt432, kdrl33, l-plastin34, mpx35, notch1b32, notch332, pu.136, runx137, gata2b38 and scl39. Embryos were then scored manually, imaged and genotyped. Separate PCR reactions were used for detecting wild-type and mutant sf3b1 alleles. A common reverse primer (5′-GACAATCACCCACGGCCATG-3′) was paired with unique forward primers for the wild-type (5′-GATCGCCAAAACACATGATG-3′) or mutant alleles (5′-CTGTCCATCTGTTCCTGAC-3′).

Flow cytometry

For generation of single cell suspensions, 10-20 embryos were first removed from their chorions using pronase (Roche), and then homogenized by manual dissociation using a sterile razor blade followed by digestion with Liberase (Roche). For the digestion, dissociated embryos were resuspended in 3 ml 1× Dulbeccos-PBS (D-PBS) (Life Technologies) supplemented with a 1:65 dilution (46 μl) of 5 mg/ml Liberase and then incubated at 37°C for 8 minutes. The reaction was s topped with 5% (157μl) fetal bovine serum (FBS) (Life Technologies). The cells were then filtered twice through 40-μm cell strainers (Falcon) and pelleted by centrifugation at 3000 rpm for 5 minutes. Cell pellets were resuspended in 1-2 ml FACS buffer (0.9× D-PBS, 5% FBS, 1% Penn/Strep (Life Technologies)). DAPI (4′,6-diamidino-2-phenylindole) was added to a final concentration of 1 μg/ml to facilitate exclusion of dead cells from the analysis. Samples were analyzed with a LSRII flow cytometer (BD Biosciences) and FlowJo version 10.0.8.

Fluorescent imaging

Embryos (24-36 hpf) were anesthetized with tricaine and oriented in a drop of 3% wt/vol methylcellulose and then mounted in 1% agarose in 35-mm imaging dishes (MatTek) as previously described40. Fluorescent imaging was performed with a Zeiss Axio Observer A1 with an AxioCam ICM1 camera and Zeiss Zen 2 software. Fluorescence was detected with GFP and Texas-Red filters.

May-Grunwald Giemsa staining of primitive erythroid cells

Two-day old embryos were placed on poly-L-lysine coated slides in a drop of 1× D-PBS + 1% bovine serum albumin (Sigma). Blood cells were released from the embryo by puncturing the pericardial sac and upper yolk sac with fine forceps. The slides were air-dried at room temperature prior to staining. For staining, slides were immersed in undiluted May-Grunwald stain (Eng Scientific May-Grunwald stain solution 1, Fisher Scientific) for 2 minutes and briefly rinsed in ddH20. Slides were then immersed in dilulted Giemsa stain (diluted 1:4 with milliQ water) for 20 minutes (Eng Scientific May-Grunwald stain solution II, Fisher Scientific) and briefly rinsed in ddH20. Once slides were dry, a drop of Permount solution (Fisher Scientific) was added and slides were covered with cover slips and left overnight to dry. Once slides were dry, the cells were visualized with a 63× oil-immersion lens.

O-dianisidine and apoptotic marker analysis

O-dianisidine staining was performed as described previously35. Dechorionated, live embryos were soaked in o-dianisidine staining solution (0.62mg/ml o-dianisidine (Sigma), 10.9 μM sodium acetate, and 0.65% H2O2) for 20 min in the dark. Acridine Orange staining was performed as previously described41. Dechorionated, live embryos were soaked in a 2μg/ml acridine orange (Sigma) solution for one hour in the dark. Immunofluorescence analysis was performed for active caspase 3 (BD Biosciences) at 24 hpf as previously described42.

Treatment with SNAP

Embryos from an incross of sf3b1 mutants were treated with the Nitric Oxide (NO) donor SNAP (S-Nitroso-N-Acetyl-D,L-Penicillamine) from 7 hpf until 28 hpf. Doses tested were based on previous studies43 and included 10μM and 30μM. Control embryos were treated with 0.06% DMSO as a vehicle control.

Statistics

For pairwise comparisons, the Student’s t-test (unpaired, two-tailed) was employed. For comparison of population distributions, Chi-squared analysis was performed. Statistical test utilized is designated in the figure legend for each analysis.

Results

Erythroid maturation is hindered in sf3b1 mutants

To characterize the blood compartments in sf3b1 zebrafish, we first assessed primitive hematopoietic induction. Primitive erythroid and myeloid cells arise from scl-positive mesoderm3,39. We observed no difference in scl expression between sf3b1 mutants and wild-type siblings at the 12 somite stage (ss)/14hpf (Figure 1A). Gata1 is an erythroid-specific transcription factor that is essential for erythropoiesis44, thus to determine if erythroid cell identity was properly specified in sf3b1 mutants, we performed flow cytometry analysis of sf3b1 Tg(gata1:eGFP) embryos at 24 hpf. We observed a subtle, but significantly higher percentage of gata1 positive cells in sf3b1 mutants compared to wild-type siblings (Figure 1B-D). At 48 hpf the number of gata1-positive hematopoietic cells remained high in mutants, but was reduced in wild-type siblings (Figure 1E). The reduction in gata1-positive cells in wild-type embryos is consistent with previous studies that showed a down-regulation of gata1 mRNA at 36-48 hpf thought to be associated with erythroid differentiation or maturation45,46. The perdurance of high gata1-expressing cells in sf3b1 mutants might be a reflection of a block in differentiation or maturation.

Figure 1. sf3b1 mutants have normal erythroid cell development but lack mature erythrocytes.

Figure 1

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A. In situ hybridization of the primitive hematopoietic marker scl at 12 ss (14 hpf) in wild type (top) and sf3b1 mutants (bottom). Number shown in the lower left corner denotes the number of embryos of each genotype that displayed a similar phenotype to the image. B. Flow cytometry analysis of gata1:eGFP cells from wild type (left) and sf3b1 mutants (right). Averages +/− standard deviation of 3 replicates are shown. For each replicate, 10-20 embryos were pooled for the analysis. C. Graph showing the percentage of gata1:eGFP positive cells shown in B. Student’s t-test **p=0.007. D,E. Schematic showing area of the embryo shown in images below. Fluorescent images of gata1:eGFP wild type (top) and sf3b1 mutants (bottom) at 24 hpf (D.) and 48 hpf (E.), scale bar: 100 μm. F. In situ hybridization of the erythroid marker β-globin at 24 hpf in wild type (top) and sf3b1 mutants (bottom). G,H. O-dianisidine staining of functional hemoglobin in mature primitive erythrocytes in wild type (top) and sf3b1 mutants (bottom) at 36 hpf (G.) and 48 hpf (H.). I. Representative images of orthochromatophilic erythroblasts stained with May Grunwald-Giemsa from wild type (left) and sf3b1 mutants (right), scale bar: 20 μm. J. Number of cells that were classified by morphology at each differentiation stage with percentages shown in brackets for wild type and sf3b1 mutants. Chi-squared test ****p=0.0001.

To investigate the differentiation status of erythroid cells in sf3b1 mutants, we examined the expression of the differentiation marker β-globin by in situ hybridization. Expression of β-globin, a component of oxygen-carrying hemoglobin, was normal in sf3b1 mutants at 24 hpf (Figure 1F). Functional hemoglobin can be detected in wild-type red blood cells beginning around 36-48 hpf using o-dianisidine staining47. In sf3b1 mutants at 36 hpf, we observed little o-dianisidine-positive erythroid cells (Figure 1G). Mutants display some developmental delay that becomes more severe as the embryos get older, thus we also looked at o-dianisidine staining at 48 hpf and noted no improvement to hemoglobin function (Figure 1H). Moreover, the level of o-dianisidine staining in the 48 hpf embryos is lower than wild types at 36 hpf indicating the decrease in o-dianisidine is unlikely solely due to developmental delay. To validate the findings in the mutants, we also tested hemoglobin function in sf3b1 morpholino-injected embryos21, and noted a similar decrease in o-dianisidine staining (Figure S1A). To further characterize the defects in primitive erythropoiesis in sf3b1 mutants, we analyzed the morphology of isolated red blood cells using May-Grunwald-Giemsa staining (Figure 1I). Mutants had a higher percentage of proerythroblasts and a corresponding decrease in more mature erythroid precursors at 48 hpf (Figure 1J). Furthermore, the later stage orthochromatophilic erythroblasts in the sf3b1 mutants showed a megaloblastoid-like phenotype compared to wild type (Figure 1I). Together, our results indicate sf3b1 mutant erythroid cells have a late-stage block in differentiation.

Myeloid cell development is impaired in sf3b1 mutants

During development, primitive myelopoiesis originates in both the anterior and posterior lateral plate mesoderm from scl-expressing cells2. As mentioned above, scl expression was unchanged in sf3b1 mutants, either in the anterior or posterior blood islands (Figure 1A). The transcription factor pu.1 is a master regulator of myeloid cell fate and is first expressed in developing zebrafish around 12 hpf/6ss36. In sf3b1 mutants, pu.1-expressing cells are detected at 22 hpf, but are slightly diminished and less dispersed across the yolk, suggesting a potential delay in differentiation (Figure 2A). We next examined the expression of markers of mature myeloid cells. Differentiation of myeloid cells is marked by expression of genes such as l-plastin or myeloperoxidase (mpx), beginning around 16 hpf and 19 hpf, respectively. At 24 hpf, expression of l-plastin and mpx are greatly diminished in sf3b1 mutants (Figure 2B,C). To rule out a contribution of developmental delay for the lack of mature myeloid marker expression in sf3b1 mutants, we examined the expression of l-plastin and mpx at 28 hpf. The markers were still diminished in the mutants, suggesting a block in myeloid cell differentiation (Figure 2D,E). We confirmed the decrease in mpx at 28 hpf in sf3b1 morpholino-injected embryos (Figure S1B). To further test this hypothesis, we used flow cytometry to quantify the frequency of lyz:dsRed myeloid cells at 36 hpf. Lyz:dsRed myeloid cells were over 22-fold decreased in sf3b1 mutants compared to wild-type siblings (Figure 2F,G). Erythroid-myeloid progenitors are also arising in zebrafish embryos at this time during development48,49. To characterize this cell population, we examined pu.1 at 26 hpf and l-plastin and mpx at 30 hpf within the posterior blood island (PBI) (Figure 3). As in the primitive wave, we observed some pu.1-positive cells in the PBI, but greatly diminished expression of more differentiated markers. Taken together, these results show myeloid specification initiates normally in the mutants, but myeloid cell differentiation is hindered.

Figure 2. Primitive myeloid cell development is impaired in sf3b1 mutants.

Figure 2

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A-E. In situ hybridization for myeloid markers. Images of wild type are shown on the left and sf3b1 mutants are shown on the right. Number shown in the lower right corner denotes the number of embryos of each genotype that displayed a similar phenotype to the image. A. The myeloid progenitor marker pu.1 at 22 hpf. B,D. The myeloid differentiation marker l-plastin at 24 hpf (B.) and 28 hpf (D.) C,E. The neutrophil marker mpx at 24 hpf (C.) and 28 hpf (E.). F. Flow cytometry analysis of lyz:dsRed-positive myeloid cells from wild type (left) and sf3b1 mutants (right) at 36 hpf. Averages +/− standard deviation of 3 replicates are shown. For each replicate, 10-20 embryos were pooled for the analysis. G. Graph showing the percentage of lyz:dsRed positive cells shown in F. Student’s t-test ****p =0.0001.

Figure 3. Erythroid-myeloid progenitor development is impaired in sf3b1 mutants.

Figure 3

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A-C. In situ hybridization for myeloid markers. Images of wild type are shown on the left and sf3b1 mutants are shown on the right. Number shown in the lower right corner denotes the number of embryos of each genotype that displayed a similar phenotype to the image. Inset to the right shows a higher magnification view of the PBI region of the embryo boxed in the image on the left. A. The myeloid progenitor marker pu.1 at 26 hpf. B. The myeloid differentiation marker l-plastin at 30 hpf. C. The neutrophil marker mpx at 30 hpf.

HSPC formation is inhibited in sf3b1 mutants

We next examined if there were changes to definitive HSPC formation. HSPCs emerge from hemogenic endothelial cells that reside along the ventral wall of the dorsal aorta3,50. One of the earliest markers of HSPCs is runx137. Expression of runx1 is nearly absent within the aorta of 28 hpf sf3b1 mutants, but is readily detected in wild-type siblings (Figure S2A). Mutant sf3b1 embryos have a variable reduction in blood flow and decrease in aortic lumenization, thus we wanted to determine if these defects were the main contributors to the observed HSPC loss in mutants. Although blood flow is needed for HSPC formation, it is dispensable for the early induction of runx1 expression at 24 hpf51. To determine if the decrease in HSPC gene expression is present at this earlier stage, we examined expression of runx1 and another HSPC marker, gata2b38, at 24 hpf. In agreement with the results at 28 hpf, we observed diminished expression of both markers at 24 hpf (Figure 4A, B). We confirmed the decrease in runx1 expression at 24 hpf in sf3b1 morpholino-injected embryos (Figure S1C). Newly-born HSPCs co-express markers of hematopoietic and endothelial cells52. This property can be visualized in embryos that are double-transgenic for c-myb:gfp and kdrl:dsred (Figure 4C). We incrossed c-myb:gfp;kdrl:dsred transgenic sf3b1 heterozygotes and quantified HSPC number at 38 hpf in mutant and wild-type siblings. Wild-type embryos had 5.4 c-myb:gfp;kdrl:dsred double positive cells on average, while sf3b1 mutants had only 1.4 (Figure 4C,D). Due to the variable decrease in blood flow and aortic lumen size observed in sf3b1 mutants, we quantified c-myb:gfp;kdrl:dsred double positive cells in embryos with different lumen sizes, but noted no significant difference in the number of double positive cells, suggesting the defect in HSPC formation is independent of these defects. To further confirm that the decreased HSPC production was not due to loss of signals from blood flow, we treated embryos with the nitric oxide donor SNAP (S-Nitroso-N-Acetyl-D,L-Penicillamine), which was previously shown to restore HSPC levels in mutants lacking blood flow43,51. Treatment with SNAP had no effect on runx1 levels in mutant embryos (Figure S2B). Combined, these data demonstrate a severe defect in HSPC formation in sf3b1 mutants independent of the effects on blood flow and aortic lumenization.

Figure 4. sf3b1 mutants have diminished HSPCs.

Figure 4

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A,B. In situ hybridization of the HSPC markers runx1 (A.) and gata2b (B.) at 24 hpf in wild type (top) and sf3b1 mutants (bottom). Inset to the right shows a higher magnification view of the AGM region of the embryo boxed in the image on the left with aorta marked with arrowheads. Number shown in the lower left corner denotes the number of embryos of each genotype that displayed a similar phenotype to the image. C. Representative fluorescent images of cmyb:gfp+;kdrl:dsred+ HSPCs (arrows) from wild type (top) and sf3b1 mutants (bottom) at 38 hpf. Middle image depicts a mutant with a less inflated aorta and the bottom image shows a mutant with a more inflated aorta. Scale bar: 50μM. Schematic on the right shows the area of the embryo shown in the fluorescent images. D. Graph showing quantification of HSPCs (cmyb:gfp+;kdrl:dsred+ cells) in wild type and sf3b1 mutants at 38 hpf. Each dot represents the number of HSPC per embryo and the line denotes the mean. Dark blue dots are HSPC counts for mutants with less inflated aorta and the light blue dots are HSPC counts for mutants with more inflated aorta. Student’s t-test ****p=6×10−11.

Aortic specification and Notch signaling are unperturbed in sf3b1 mutants

Hemogenic endothelial cells in the ventral wall of the dorsal aorta are the immediate precursor to HSPCs, thus proper vasculature formation is an essential prerequisite for HSPC induction50. We next analyzed the expression pattern of endothelial markers in sf3b1 mutants and wild-type siblings. The pan-endothelial marker kdrl is expressed in the dorsal aorta, cardinal vein, and intersomitic vessels in both wild-type and mutant siblings at 24 and 28 hpf (Figure 5A and S3A). Proper artery-vein identity precedes HSPC induction3,50. To determine if vessel identity was correctly initiated in sf3b1 mutants, we examined artery and vein-specific markers. The vein marker flt4 shows the presence of a well-formed posterior cardinal vein in 24 and 28 hpf mutant embryos (Figure 5B and S3B). Aorta-specific markers ephrinb2, notch1b, and notch3 are expressed in the aorta of both wild type and mutants at 24 and 28 hpf, demonstrating proper specification of aortic endothelium (Figure 5C and S3C). The Notch pathway gives arterial identity to endothelial cells, and is a critical signal for HSPC specification29,50. Correct expression of notch1b and notch3 within the dorsal aorta suggests an intact signaling network, but does not test if the pathway is active. Notch pathway activation can be visualized using tp1:eGFP Notch reporter fish29. At 36 hpf, mutant embryos show the same pattern of Notch signaling within the aorta and intersegmental vessels compared to their wild-type siblings (Figure 5D). These data indicate the lack of HSPC formation is not due to a lack of Notch signaling within arterial cells.

Figure 5. Endothelial markers are expressed in the correct vessels in sf3b1 mutants.

Figure 5

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A-C. In situ hybridization for vascular markers at 24 hpf. Images of wild type are shown on the left and sf3b1 mutants are shown on the right. Number shown in the lower right corner denotes the number of embryos of each genotype that displayed a similar phenotype to the image. Inset to the right shows a higher magnification view of the AGM region of the embryo boxed in the image on the left with aorta or vein marked with arrowheads. A. The pan-endothelial marker kdrl. B. The vein marker flt4. C. The aorta-specific markers ephrinb2, notch1b and notch3. D. Schematic showing area of the embryo shown in images below. Representative fluorescent images of tp1:eGFP (Notch reporter) wild type (left) and sf3b1 mutants (right) at 36 hpf. Arrowhead denotes the aorta.

The results suggest that the hematopoietic defects in sf3b1 mutants are likely due to problems transitioning between different cellular states, but cell death cannot be excluded as a contributing factor. We assessed apoptosis in sf3b1 mutants using acridine orange staining and antibody staining for active caspase-341,42. While a clear increase in cell death in neuronal tissues, such as the brain and spinal cord, was observed, we noted no significant increase in apoptosis in other tissues in sf3b1 mutants compared to wild-type siblings (Figure S4). These results support the model that splicing controls hematopoietic differentiation decisions.

Discussion

Recent clinical findings and discoveries in zebrafish development indicate that normal functioning of the spliceosome is critical for hematopoiesis. These studies also implied that the ensuing hematopoietic defects were somewhat distinct depending on the spliceosomal component affected. The most commonly mutated spliceosomal factor in MDS is SF3B1, yet there is currently no in vivo model describing what aspects of hematopoiesis are affected when this factor is lost. Here we demonstrated that depletion of sf3b1 during zebrafish development results in deficits in primitive and definitive hematopoiesis. Primitive erythroid cells in sf3b1 mutants arise and initially differentiate normally, but then arrest at the orthochromatophilic stage and take on an abnormal megaloblastoid-like morphology. Primitive myelopoiesis also initiates normally, but then pu.1-expressing progenitors fail to mature. These data indicate sf3b1 is needed for maturation of myeloid and erythroid progenitors. In contrast, induction of definitive hematopoiesis is severely impaired. Expression of the hemogenic endothelial and HSPC markers runx1 and gata2b is hampered as early as 24 hpf although the specification of non-hemogenic aortic endothelial cells at the same time and place in development appears mostly unaltered. These data suggest that sf3b1 is differentially required in distinct endothelial cell populations and that hematopoiesis is acutely sensitive to perturbations in the spliceosome.

A common feature among the hematopoietic defects in sf3b1 mutants is inhibited differentiation and maturation. Recent RNA-seq studies have revealed a complex orchestration of splicing throughout erythroid and myeloid differentiation53-57. The functional importance of these splicing events remains unclear. In erythroid maturation, splicing changes are greatest in the later stages of differentiation55,56. Our data provide a functional corollary to the importance of this observation as erythroid differentiation in sf3b1 mutants is blocked at a late progenitor stage. Unlike erythroid and myeloid splicing patterns, the splicing events occurring during the endothelial-to-hematopoietic transition are unknown. The lack of induction of expression of the key hemogenic transcription factors runx1 and gata2b demonstrate splicing is a regulator of definitive HSPC origins. Further exploration of the splicing aberrations in sf3b1 endothelial cells will help decipher which specific splice isoforms are important in HSPC development.

Although many spliceosomal mutants characterized to date have hematopoietic perturbations, they are not entirely overlapping, even in factors that are within the same complex. U2 snRNP is a large macromolecular complex made up of nearly 20 factors. Zebrafish mutants of three of these factors (sf3a315, ddx46/prp512, and now sf3b1) have been characterized, and all display defects in the hematopoietic system. Mutants in sf3a3 have normal primitive erythropoiesis and myelopoiesis, but show defects in HSPC formation, while ddx46 mutants initiate HSPC induction normally, but have problems maintaining their stem cell pool. These studies lead to two non-mutually exclusive hypotheses for the differences observed: 1) different splicing events are affected in each spliceosomal mutants and 2) the diverse phenotypes could be due to non-spliceosomal functions of each splicing factor. Indeed, recent studies have found that SF3B1 has some non-splicing functions in responding to various cellular stressors58,59.

These differences can also be observed clinically. MDS is a pleiotropic hematopoietic disease marked by perturbations to the erythroid and myeloid lineages. U2AF1 is an essential factor that associates with the U2 snRNP complex to help recruit it to splice sites. Despite the interconnection between these two complexes in splicing, mutations in U2AF1 and SF3B1 are associated with different features of MDS. Mutations in U2AF1 segregate with MDS with poor overall survival, while mutations in SF3B1 show a strong correlation with refractory anemia with ringed sideroblasts (RARS) and an overall favorable prognosis4-10. Erythroid cells in RARS are poorly differentiated and show defects in iron usage resulting in hemoglobinization problems. Reminiscent of this type of MDS, zebrafish sf3b1 mutants have poorly differentiated red blood cells and low levels of functional hemoglobin, but they do not show symptoms of myeloproliferation associated with more malignant forms of MDS. Whether these differences are due to differences in splicing abnormalities or in non-splicing functions of U2AF1 and SF3B1 requires further exploration.

Combined, the clinical indications and differences in developmental hematopoietic deficiencies among spliceosomal mutants show that there is still much to learn about spliceosome protein function in metazoans. The zebrafish is an excellent model for addressing this problem. Maternal contribution of essential factors like spliceosomal components allows survival of embryos to later stages during organogenesis permitting comparison across many tissues. Zebrafish are highly amendable to genetic and chemical perturbations, which can be employed to define the functionally relevant pathways for each tissue type affected. Genome-wide studies of blood diseases, such as MDS, have revealed a number of correlations between gene mutation and disease phenotype, which are difficult to dissect due to the complexity of the mutational landscape within the heterogenous mix of cells analyzed. The use of in vivo systems, such as the zebrafish, provides a facile genetic approach for testing the contribution of single gene perturbations. We have now established numerous phenotypic similarities between zebrafish sf3b1 mutants and SF3B1-mutated MDS. Future studies will take advantage of this model to uncover the molecular and cellular origins of the hematopoietic defects arising from deficiencies in sf3b1.

Supplementary Material

01

Highlights.

  • Erythroid and myeloid differentiation is inhibited in sf3b1 zebrafish mutants.

  • Sf3b1 is required for hematopoietic stem and progenitor cell emergence (HSPC).

  • Loss of HSPCs is independent to defects in endothelial cell specification.

  • Notch signaling in the aorta does not require Sf3b1.

Acknowledgements

This work was funded by Aplastic Anemia and MDS International Foundation (RCC), Training Program in Cellular and Molecular Biology and Genetics, T32 GM007491 (SN), Gabrielle’s Angel Foundation, NIH 7K01DK085270-05, and NIH 6R03DK102975-02 (TVB).

We would like to thank Eirini Trompouki for critical reading of the work as well as Trista North, Leonard Zon and David Traver for sharing zebrafish strains and reagents.

Footnotes

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Authorship Contributions

ADG, RCC, SN, SP, and TVB performed experiments and analyzed data. ADG, RCC, and TVB wrote and edited the manuscript.

The authors have no conflicts to disclose.

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