1 Institute of Molecular Medicine and Genetics, Medical College of Georgia, Augusta, GA 30912. USA
2 Department of Molecular, Cell and Developmental Biology, University of California at Los Angeles, Los Angeles, CA 90095, USA
3 Howard Hughes Medical Institute, Childrens Hospital of Boston, 320 Longwood Avenue, Enders 750, Boston, MA 02115, USA
*Author for correspondence (e-mail: shuolin{at}ucla.edu)
Accepted 12 June 2002
![]() |
SUMMARY |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Zebrafish, Erythroid protein 4.1, Hereditary elliptocytosis, Congenital anemia, merlot, chablis, Marginal band
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Erythrocyte protein 4.1 (band 4.1 or 4.1R) is a multifunctional structural protein in the red cell membrane skeleton whose interaction with both transmembrane and cytoskeletal proteins plays an indispensable role in maintaining red cell morphology, membrane deformability and mechanical stability (Yawata et al., 1997; Conboy, 1993
). Human protein 4.1 was first identified in the erythroid cell membrane skeleton (Conboy et al., 1986a
), and contains four distinct structural and functional domains with molecular weights of 30, 16, 10 and 22/24 kDa (Leto and Marchesi, 1984
). Through its spectrin-binding domain at the 10 kDa domain (Correas et al., 1986
), protein 4.1R accelerates and stabilizes the interaction between spectrin and actin filaments (Ohanian et al., 1984
). This ternary complex is essential for maintaining the red cells mechanical stability (Lorenzo et al., 1994
). Red cells that are completely deficient in 4.1R or lack the spectrin-binding domain of 4.1R have abnormal elliptical cell morphology and fragile membrane (Tchernia et al., 1981
; Marchesi et al., 1990
). Protein 4.1R also plays a crucial role in anchoring the spectrin-actin framework to the overlaying lipid membrane. Via its N-terminal 30 kDa membrane-binding domain, protein 4.1R interacts with the cytoplasmic domains of the membrane proteins band 3 (Pasternack et al., 1985
), glycophorin C (Hemming et al., 1995
; Marfatia et al., 1995
), p55 (Chang and Low, 2001
; Alloisio et al., 1993
; Nunomura et al., 2000
) and CD44 (Nunomura et al., 1997
). Calmodulin also binds to this domain and modulates protein 4.1R interactions with its binding partners (Tanaka et al., 1991
; Lombardo and Low, 1994
; Nunomura et al., 2000
). The presence of the evolutionarily conserved N-terminal membrane-binding domain (also called the FERM domain: 4.1-Ezrin-Radixin-Moesin) (Chishti et al., 1998
; Hoover and Bryant, 2000
; Arpin et al., 1994
) is a common feature of a diverse group of band 4.1 related proteins such as 4.1G, 4.1N and 4.1B in vertebrates (Peters et al., 1998
), and the putative 4.1R homolog in Drosophila, also known as coracle (Fehon et al., 1994
).
Although the process of erythropoiesis in mammals and nonmammalian vertebrates is essentially the same, there are structural and morphological differences between mature red cells. The erythroid precursors of nonmammalian vertebrates, similar to their counterparts in mammals, are spherical and contain a round nucleus with open chromatin and basophilic cytoplasm. However, terminally differentiated and mature red cells of other vertebrates are nucleated and elliptical, while mammals have anucleated red blood cells with round, biconcave morphology. During differentiation, the cytoskeleton of red cells undergoes extensive reorganization that results in their final morphology (Wickrema et al., 1994). The elliptical morphology of red cells of nonmammalian vertebrates is due to their structure of cytoskeletal system, which in addition to the membrane skeleton contains intermediate filaments and an enveloping layer of microtubules called the marginal band (Cohen, 1991
; Cohen et al., 1998
). The extent of interaction between the marginal band and the protein network of membrane skeleton in red blood cells is yet to be characterized.
The availability of induced mutations that affect different aspects of hematopoiesis is one of the advantages of the zebrafish system. The zebrafish mutants merlot and chablis are members of a group of zebrafish mutants that have normal onset of primitive hematopoiesis followed by a decrease in the number of circulating red blood cells at subsequent stages of larval development (Ransom et al., 1996). One member of this group, riesling, has been characterized as having a mutation in ß-spectrin (Liao et al., 2000
).
In this work, we present the characterization of merlot/chablis phenotype and identification of zebrafish protein 4.1R as the mutated gene in merlot and chablis. We also present merlot/chablis as genetic models to study the role of protein 4.1R in morphogenesis and terminal maturation of nucleated red blood cells.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Genetic mapping and linkage analysis
A polymorphic mapping strain was generated by crossing heterozygote mottu275 with the polymorphic WIK strain. Embryos were collected from pair mating of Tü/WIK heterozygotes and phenotyped at 96 hours post fertilization (hpf) for anemia. Genomic DNA extraction from individual embryos and bulk segregant analysis (BSA) was performed as described (Talbot and Schier, 1999; Zhang et al., 1998
). Simple sequence-length polymorphism (SSLP) markers (Shimoda et al., 1999
; Knapik et al., 1998
) used for BSA were selected from the MGH web server (http://zebrafish.mgh.harvard.edu) and purchased from Research Genetics (Huntsville, AL). Diploid mutant cha and wild-type embryos were collected from AB/Darjeeling and AB/SJD heterozygotes. Genome-wide scanning on BSA pooled DNA was used for linkage analysis. Close microsatellite markers, z11376 and z13511, were found to flank the cha locus.
Selection of candidate gene and isolation of cDNA
Single wild-type and mutant embryos were genotyped with SSLP markers z25218, z10036 and z25278, and the genetic locus defined by them was searched for cloned ESTs (WUZGR, http://zfish.wustl.edu). Two cDNA clones, Fc37c08 and Fb70c02, were identified as potential candidate genes and were purchased from Incyte Genomics (St Louis, MO). Fc37c08 was a partial clone of 2 kb and Fb70c02 was a full-length 5 kb sequence of the cDNA encoding for a protein homologous to human erythroid protein 4.1. Total RNA from adult tissue (blood and kidney) and single wild-type and mutant (both mot alleles) embryos was extracted by Trizol (Gibco-BRL, Rockville, MD). The first strand cDNA was synthesized using the SuperScript First-Strand Synthesis System for RT-PCR (Gibco-BRL) following the manufacturers protocol. The oligo (dT) primed first strand cDNA was subjected to PCR amplification using the Expand Long Template PCR System, (Roche, Indianapolis, IN) with a 5'- GATCATTGCCGGACATGTAAA-3' forward primer and a 5'-TGTAAGCGGGTGAAATAAGCT-3' reverse primer. The amplified full-length 5Kb cDNA fragment of the zebrafish erythroid protein 4.1R was cloned into the PCR 2.1 TOPO TA vector (Invitrogen, Carlsbad, CA) and sequenced (primer sequences available upon request).
The cha locus was independently mapped near SSLP markers, z11376 and z13511, which allowed the identification of a candidate EST, Fb70c02, within this genetic interval. A large P1 artificial chromosome (PAC) clone, 215:J16, was isolated by hybridization with the Fb70c02 EST clone. The termini of PAC 215:J16 were sequenced to derive simple sequence conformational polymorphic (SSCP) primers (forward, 5'-TCGTCGCTCAGTCATCAAGTAACA- 3'; reverse, 5'-TGACATGAACCTTCGCTTCCC-3') for analyzing embryos that were genetic recombinants with markers, z11376 and z13511.
Mutation and linkage analysis using allele-specific primers
To confirm the mutation, RT-PCR on total RNA from several 24 hpf mutant embryos was performed and the PCR products were sequenced. For the mottu275 allele, a forward primer (P10; 5'-GTGACAACAGAGGAGATCCAA-3') and a reverse primer (P14; 5'- AACACCTCAACAGCCGAACC-3') were used to amplify a 630 nucleotide fragment that contained the point mutation. For mottm303c, the forward primer P10 was used with a reverse primer (P4; 5'- CGATTCAGCCTTTTCTCTCTA-3') to amplify a 1.4 kb fragment containing the point mutation. Linkage analysis with allele-specific primers (Bottema and Sommer, 1993; Newton et al., 1989
) was used to confirm the linkage of the anemic phenotype to the genomic sequence. PCR was performed on genomic DNA extracted from adult heterozygotes and 96 hpf mutant (mottu275) and wild-type embryos. A common forward primer (P10) was used with wild-type-specific reverse primer (WtR; 5'-ACCTCAACAGCTGGACCTCG-3') or mottu275-specific reverse primer (MutR; 5'-ACCTCAACAGCTGGACCTCA-3') (point mutation G to A is underlined) to amplify
550 bp fragment of genomic DNA. PCR conditions were optimized so that the primer pair P10 and WtR only amplified wild-type DNA, whereas the primer pair P10 and MutR only amplified mutant DNA. For confirmation of the chatu242e and chatu245 nonsense mutation, genomic DNA was amplified with primers (forward, 5'-GATGTGGAGGACGACTGGTTTATC-3'; reverse, 5'-CTTCGTCTCTGGTACTGTTATCTGTTC-3'). The PCR product was analyzed by allele-specific oligonucleotide hybridization (Farr et al., 1988
) with either wild-type (5'-TTTTTAGTGCRAGGTCCRG-3') or mutant (5'-TTTTTAGTGTRAGGTCCRG-3') oligonucleotide.
Truncation mutation analysis
The full-length 5 kb cDNA fragments from wild-type and mutant (mottu275) fish were cloned into PCR 2.1 TOPO TA vector (Invitrogen) and were subjected to in vitro transcription and translation using the TNT T7 reticulocyte lysate system (Promega, Madison, WI). Synthesized wild-type and mutant proteins were labeled by including [35S] methionine in the reaction, and were resolved by a 10% SDS-PAGE, which was dried and exposed to autoradiographic film.
In situ hybridization
Whole-mount in situ hybridization with digoxigenin-labeled RNA probes was performed on 24 hpf wild-type and mutant embryos (both mot alleles) as described (Westerfield, 1993) with some modifications. A fragment of zebrafish P4.1R cDNA (from 97 to +1575 bp) was amplified by RT-PCR from wild-type adult blood, subcloned into PCRII vector (Invitrogen), and used as a template for generating sense and antisense RNA probes. The full-length cDNA clone, Fb70c02, was also used as a template for probe synthesis. The construct was digested by KpnI and the RNA probe spanning the 3'UTR and poly-A tail was synthesized using SP6 RNA polymerase. A probe for gata1 was generated as described elsewhere (Long et al., 1997
). cDNAs encoding for embryonic globins were isolated from our embryonic blood-specific cDNA library. Briefly, inserts were amplified by PCR using flanking vector primers containing T7 RNA polymerase site. The PCR products were used as templates to generate probes.
Blood analysis, tissue preparation and o-dianisidine staining
Adult fish were anesthetized in a solution of 0.05% tricaine (Sigma) and peripheral blood was drawn by cardiac puncture using heparinized pulled micro needles. Embryonic blood was collected by cutting the tail of embryos in a PBS solution containing tricaine, sodium citrate and albumin. Collected embryonic blood cells were spun onto a glass slide at 750 rpm for 3 minutes using a Cytospin 2 (Shandon, Pittsburgh, PA). Blood smears were stained with Wright-Giemsa method following the manufacturers instruction (Sigma). The red blood cell number of wild-type and mutant adult fish was determined using a Neubauer hemocytometer. Hemoglobin was measured using Drabkins reagent (Sigma). Exactly 2 µl of blood was suspended in 1 ml of Drabkins reagent and incubated at room temperature for 10 minutes. The optical density was then measured at a wavelength of 540 nm using a Smart Spec 3000 spectrophotometer (BioRad, Hercules, CA). Red blood cell indices were determined using a GEN®S Coulter Counter (Miami, FL). Kidney and spleen smears from adult wild-type and mutant fish were prepared as described (Long et al., 2000). Analysis of hemoglobin expression in embryos was performed using o-dianisidine, as described previously (Detrich et al., 1995
).
Osmotic fragility
Freshly drawn blood from wild-type and homozygous adult fish was washed three times in 0.9% NaCl and suspended in saline to 10% packed cell volume. Cell suspension (5 µl) was added to each of 150 µl solutions that contained 0.900%, 0.800%, 0.700%, 0.600%, 0.500%, 0.475%, 0.450%, 0.425%, 0.400%, 0.300% and 0.200% of NaCl (wt/vol). Cells were incubated at room temperature for 15 minutes, and then centrifuged for 3 minutes at 420 g. The percent hemolysis in the supernatant was measured at 540 nm with the spectrophotometer.
Electron microscopy and TUNEL assay
For scanning electron microscopy, blood was drawn from adult wild-type and homozygous mutant fish and washed three times in cold saline. Blood cells were crosslinked in buffered glutaraldehyde and post fixed in osmium tetroxide. Dehydrated cells were rinsed in hexamethyldisilazane, sputter-coated with gold and examined by a Cambridge 360 Scanning Electron Microscope (Leo Electron Microscopy, Cambridge, UK). For transmission electron microscopy, freshly drawn blood cells from wild-type and mot fish were crosslinked and dehydrated as above, embedded in Epon, and 60 nm sections stained with uranyl acetate and lead citrate were examined with a JEOL JEM-100CX electron microscope (JEOL, Ltd. Tokyo, Japan). ApopTag (Intergen, NY) kit was used for detection of apoptotic cells in peripheral blood, kidney and spleen.
Immunofluorescence and confocal microscopy
Blood smears from adult wild-type and homozygous mot fish were prepared as described above. Cells were extracted in 0.2% Triton X-100 in phosphate-buffered saline (PBS) for 30 minutes, fixed in 4% paraformaldehyde for 20 minutes at room temperature, and then post fixed in methanol for 10 minutes at 20°C. Cells were incubated in 1% bovine serum albumin (BSA)/PBS for 1 hour, incubated with anti-tubulin monoclonal antibody (Sigma) in 1%BSA/PBS for 1 hour, washed three times in PBS, incubated with Cye3-conjugated anti-mouse IgG antibody (Jackson ImmunoResearch Laboratories, West Grove, PA) for 1 hour, washed 3 times in PBS and counterstained with DAPI.
Rescue experiment and microinjection
A zebrafish protein 4.1-GFP fusion construct was generated by RT-PCR. Two primers, rescF (5'-AAGCTTCCCGCTTTTGCAGATCA-3') and rescR (5'-GGTACCTCGAGCAAATATTTCT-3'), were used to amplify the full-length P4.1 cDNA and remove the stop codon. The PCR product was cloned in frame into the green fluorescent protein (GFP) expression vector pEGFP-N3 (Clontech). The integrity of the construct was verified by restriction mapping and sequencing. Fertilized eggs from a cross of mottu275 homozygotes were dechorionated and injected at the one- to two-cell stage with P4.1-GFP construct (150 ng/µl) as described (Meng et al., 1999). Injected embryos were incubated at 28°C, examined for GFP expression, phenotyped and stained with o-dianisidine at 96 hpf.
GenBank Accession number
AY124488 for full-length zebrafish P4.1R cDNA and protein sequence.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Hematological and pathological analysis of adult mot and cha fish
About 5-10% of homozygous embryos could be raised to adulthood with frequent feeding and ample supply of oxygenated water. mot adult fish suffer from growth retardation and appear pale. Moreover, cardiomegaly is observed in most adult fish, predominantly in males (Fig. 2). Analysis of peripheral blood from adult homozygotes showed evidence of severe anemia. Red blood cell count and hemoglobin levels were significantly decreased in mot fish (Table 1). Despite a severe reduction in the number of red cells and hemoglobin concentration, adult mot are viable, fertile and appear to exhibit normal fish behavior. Examination of mot peripheral blood revealed microcytic cells that exhibit differentiation arrest at the basophilic erythroblast stage with severe abnormal membrane morphology, as seen in Fig. 2A. About 1% binucleated cells were also detected in blood smears of mot fish, whereas in wild type no binucleated cells were observed (data not shown). Examination of stained kidney smears revealed a marked increased in the number of erythroid progenitors in mot mutant (Fig. 2B), indicating a reactive hematopoiesis. Evidence of membrane abnormalities was also noted in a few proerythroblasts, suggesting that abnormal membrane structure may have an impact in the early stages of erythroid differentiation. When adult mot fish were dissected (Fig. 2D), prominent features included enlarged gall bladder and spleen, as well as an increased concentration of bilirubin in the internal organs (liver, GI tract). Hypercellular and enlarged kidneys (the site of definitive hematopoiesis in adult fish) were also observed in homozygous fish, resulting from a compensatory mechanism for excess hemolysis and reduced survival of erythroid cells.
|
|
These data indicate that mot fish suffer from a severe and partially compensated hemolytic anemia with symptoms similar to hemolytic anemia in humans. The enlarged gall bladder is especially intriguing because such enlargement usually occurs in humans in many cases of hereditary anemia because of obstructive bilirubin gallstones.
Osmotic fragility test is a sensitive test for detection of cells with altered tolerance to osmotically induced stress. Although in humans increased osmotic fragility is characteristic of hereditary spherocytosis, we hypothesized that as the membrane structure of red blood cells in mot fish is abnormal, as seen on peripheral blood smears, the osmotic fragility test may be informative. Our results indicated that mot cells were, indeed, profoundly susceptible to osmotic stress (Fig. 3). The salt concentration in which 50% of wild-type cells undergo hemolysis is significantly lower (0.425%) compared with mutant cells (0.700%). This means that, compared with wild-type cells, mot cells are considerably more fragile in hypotonic solutions.
|
|
We attempted to investigate whether the formation of marginal band of microtubules is compromised in mot cells. Immunohistochemistry analysis showed (Fig. 4E) that in wild-type red blood cells microtubules are confined to a marginal band at the long axis of cells. By contrast, mot cells exhibit a diffuse pattern of microtubule filaments in different planes of the cell, indicating that the absence of protein 4.1 in the red cell membrane cytoskeleton affects the proper assembly of microtubules into a marginal band at the periphery of red blood cells.
Based on phenotypical, hematological and molecular analysis of the zebrafish mot, we hypothesized that the defect in mot gene is intrinsic to red cells, and that the mot is a structural component of erythroid cell membrane. Based on this hypothesis and the results of positional cloning experiments, we searched the available cloned and mapped EST databases for possible candidate genes.
Positional cloning of mot and cha with a candidate gene approach
We used bulk segregant analysis to map the mot gene on linkage 16. Because the available genetic markers flanked a sizable physical distance, chromosomal walking was not feasible and we decided to approach the problem by examining cloned ESTs. As we had already hypothesized that the mutation in mot is likely to affect red blood cells membrane integrity, it was fortunate that we located the EST encoding the zebrafish protein 4.1. DNA sequence analysis from wild-type and both mottu275 and mottm303c alleles, revealed nonsense point mutations in the mot gene causing premature stop codons that would cause a truncated protein 4.1. In mottu275 we detected a C-to-T transition, whereas in mottm303c a G-to-A transition caused premature termination codons at +2349 and +3036 nucleotides, respectively (Fig. 5A). Both mutations occurred in a GC-rich region of the cDNA, and are upstream from the spectrin-binding domain (SBD) of P4.1R, generating a truncated and dysfunctional protein that lacks the SBD. In order to confirm that point mutation would result in premature translation termination, we performed a protein truncation test using cloned wild-type and mottu275 cDNAs as templates, which detected a truncated protein product from the mutant allele (Fig. 5B). We repeated the RT-PCR and sequencing on several mutant embryos from different batches to confirm our results. In order to determine if the point mutation is linked to the mutant phenotype, we performed linkage analysis by allele-specific PCR on genomic DNA extracted from heterozygous adult, wild-type and mottu275 embryos. PCR analysis of 100 embryos did not show any recombination, indicating a close linkage between the point mutation in 4.1R and mottu275 phenotype. A representation of PCR products is shown in Fig. 5C.
|
In order to examine the spatial expression pattern of P4.1R, we used several riboprobes generated from different domains of P4.1 to perform whole-mount in situ hybridization. The results revealed that mot gene expression is erythroid-specific and restricted to the intermediate cell mass (Fig. 5F). In both alleles of mot (data not shown for mottm303c), a drastically decreased level of P4.1R transcripts was detected, which is possibly due to nonsense-mediated mRNA decay (Culbertson, 1999; Ruiz-Echevarria et al., 1998
).
We examined the level of two proteins that P4.1R interacts, band 3 and ß-spectrin, in erythroid lysates by western analysis from mot, mot heterozygous and wild-type siblings; no differences were detected in the level of band 3 and ß-spectrin protein (data not shown).
The cloned protein 4.1 cDNA contains multiple open reading frames with the longest containing 4605 nucleotides, encoding a 1535-amino acid protein with a calculated molecular weight of 180 kDa. The deduced amino acid sequence of zebrafish P4.1R derived from the nucleotide sequence was subjected to a homology search using the BLAST program at NCBI. We used ExPASY proteomics tools (http://ca.expasy.org/) to analyze zebrafish P4.1R for conserved domains and prediction of protein-sorting signals and localization sites, which revealed several putative phosphorylation and glycosylation sites along with several bipartite nuclear localization signals. Amino acid sequence comparison of zebrafish P4.1R with human P4.1R showed an identity of 58% (73% similarity) in the N-terminal FERM domain and a 41% identity (54% similarity) in the SBD (data not shown). Structural domains of zebrafish protein 4.1 include an N-terminal membrane binding domain of 330 amino acids,
1000 amino acids with a novel sequence, the spectrin binding domain of 100 amino acids, followed by 22 amino acids at the C terminus (Fig. 6). The organization of structural domains of zebrafish protein 4.1 is different from those of mammalian protein 4.1. In mammalian 4.1R, there is a short 16 kDa domain that is flanked by the FERM domain and the spectrin-binding domain. In zebrafish 4.1R, this domain is composed of approximately 1000 amino acids. We searched various databases for known homology to this region, but it appeared that this was a novel sequence with no known homologous sequence. One interesting aspect of this domain is the presence of a sequence of about 90 amino acids that repeats four times in a tandem fashion, starting at position 780 and ending at 1105. A BLAST search did not detect any known peptides with significant homology with this domain.
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The mot and cha mutations were originally thought to be different based on genetic non-complementation data (Ransom et al., 1996). However, the finding that mot and cha are allelic and encode the same genetic defect in our present study most likely reflects an error in the previous complementation data. An alternative possibility for the same mutation for both cha alleles and mottu275 would be mutability hot spot of this locus.
Although hereditary elliptocytosis caused by dysfunctional or lack of 4.1R has been reported in mammals, this is the first report in a non-mammalian vertebrate animal model. Human mutations in erythroid-specific protein 4.1R generate two groups of molecular abnormalities. The first group causes partial or complete deficiency of P4.1 in red cells, whereas the second group disrupts the erythroid-specific spectrin-binding domain of P4.1 (Lorenzo et al., 1994; Conboy et al., 1993
; Alloisio et al., 1981
; McGuire et al., 1988
; Conboy et al., 1986b
; Alloisio et al., 1982
; Tchernia et al., 1981
). The pathological consequences of both defects are the presence of mild to severe anemia known as hereditary elliptocytosis (HE). The molecular defect, as well as pathological symptoms in the zebrafish mot is similar to hereditary elliptocytosis (HE) in mice and humans. Human red cells deficient in protein 4.1R are characterized by abnormal elliptical morphology, reduced red cell deformability, increased red cell membrane fragmentation and disrupted skeletal network (Tchernia et al., 1981
; Yawata et al., 1997
). Analysis of peripheral blood showed abnormal membrane spiculation and fragmentation in mot cells, indicative of cytoskeletal fragility and membrane loss. This progressive loss of redundant membrane in mot cells results in microcytic and spherocytic cells, and generates fragile red cells with a decreased surface area to volume ratio.
Recently, a mouse knockout of protein 4.1R was generated to study the role of 4.1R in erythroid and nonerythroid cells (Shi et al., 1999). Protein 4.1 null homozygote mice suffer from a moderate hemolytic anemia, with red blood cells exhibiting abnormal morphology and increased membrane fragility. These mice are viable and reach adulthood, which is in contrast to mot/cha fish, where only 5-10% of homozygote embryos survive to adulthood. Interestingly, a majority of hematopoietic mutations in zebrafish are embryonic lethal and are characterized by a mild to severe reduction in the number of circulating red cells at 2-4 days of development (Ransom et al., 1996
). It appears that abnormal embryonic red cells in zebrafish mutants undergo an accelerated clearance from circulation before the onset of definitive hematopoiesis. The circulation in zebrafish starts shortly after 24 hpf and exposes red cells to the mechanical and chemical stress of microcirculation, which may be efficient enough to sensitize abnormal cells to early destruction.
Examination of peripheral blood from adult mot fish demonstrated morphologically abnormal cells with an apparent differentiation arrest at the late erythroblast stage. We propose two hypotheses that may explain these findings. Icteric internal organs and the enlarged spleen in adult mot fish are indicative of a severe extravascular hemolysis. Presence of immature red cells in the peripheral blood may be a consequence of premature release of basophilic erythroblasts from the kidneys in response to increased cell destruction of more mature cells. The assembly of protein 4.1 into the membrane cytoskeleton for the final stabilization of the red cell membrane skeleton occurs late during erythroid maturation (Lazarides and Woods, 1989; Lazarides, 1987
). As normal function and survival of circulating mature red cells depend on their cellular deformability and structural integrity (Weed, 1970
), the more mature red cells with accumulated membrane lesions are selectively sequestered by the spleen. This results in a left shift in erythroid maturation, which is seen as a maturation arrest in the peripheral blood. Another hypothesis is the arrest in erythroid maturation may be related to ineffective erythropoiesis. Cytological examination of the kidneys revealed a profound erythroid hyperplasia. However, the number of immature erythroid progenitors was much higher than more mature cells, suggestive of a maturation arrest and ineffective erythropoiesis. The presence of apoptotic erythroid cells in the kidneys of mot fish is consistent with recent data describing the correlation of apoptosis of erythroid progenitors and ineffective erythropoiesis in several human anemias (Pootrakul et al., 2000
; Mathias et al., 2000
). Further studies may provide insights into the role of protein 4.1R in maturation of erythroid cells in zebrafish.
Role of zebrafish protein 4.1 in erythroid morphogenesis
Terminal maturation of erythroid cells involves the development of anucleated erythrocyte from nucleated proerythroblast and is associated with dynamic changes in the membrane and cytoskeletal organization and composition (Woods and Lazarides, 1988; Bennett, 1985
; Wickrema et al., 1994
; Woods et al., 1986
). The correlations of abnormal red cell morphology with genetic defects in membrane cytoskeletal proteins are well established (Delaunay et al., 1996
; Davies and Lux, 1989
; Palek, 1987
; Mohandas and Gascard, 1999
). In non-mammalian vertebrates, terminal maturation entails the genesis of an ellipsoid nucleated red cell from its spheroid precursor. Extensive in vitro analyses of the red cells of non-mammalian vertebrates show that their elliptical morphology is partly due to the presence of marginal band of microtubules (Winckler and Solomon, 1991
; Cohen et al., 1998
; Cohen, 1991
). However, owing to the lack of a genetic model to analyze and elucidate the morphogenic mechanism of vertebrate red cells in vivo, the extent and nature of interactions between the marginal band and other cytoskeletal proteins and their possible role in erythroid maturation and morphogenesis is not defined. The profound abnormal morphology and membrane instability of mot cells, and evidence of an abnormal formation of the marginal band support the crucial role of zebrafish protein 4.1R in maintaining the integrity of the red blood cell membrane. Characterization of hereditary spherocytosis in zebrafish riesling, resulting from defective erythroid ß spectrin, also demonstrated that the aggregation of microtubule filaments into a marginal band is compromised in mutant red cells (Liao et al., 2000
). These results, along with evidence of the interaction of the marginal band with ERM proteins and F-actin (Correas et al., 1986
; Birgbauer and Solomon, 1989
), provide strong evidence that multifunctional protein elements of the membrane skeleton that form multi-protein complexes are key components in determining and sustaining the nucleated red cell morphology.
In this report, we have presented the zebrafish mutant mot as a genetic model for hereditary anemia because of the abnormal structure of the erythroid specific protein 4.1R. Over the past few years, characterization of zebrafish mutants with defects in hematopoiesis has established the zebrafish as a useful genetic model to study hematopoiesis in higher vertebrates. The zebrafish mutant merlot provides an excellent animal model with which to explore protein 4.1R structure and function further in nucleated erythroid cells.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Alloisio, N., Dorleac, E., Girot, R. and Delaunay, J. (1981). Analysis of the red cell membrane in a family with hereditary elliptocytosistotal or partial of protein 4.1. Hum. Genet. 59, 68-71.[Medline]
Alloisio, N., Dorleac, E., Delaunay, J., Girot, R., Galand, C. and Boivin, P. (1982). A shortened variant of red cell membrane protein 4.1. Blood 60, 265-267.[Abstract]
Alloisio, N., Dalla, V. N., Rana, A., Andrabi, K., Texier, P., Gilsanz, F., Cartron, J. P., Delaunay, J. and Chishti, A. H. (1993). Evidence that red blood cell protein p55 may participate in the skeleton-membrane linkage that involves protein 4.1 and glycophorin C. Blood 82, 1323-1327.[Abstract]
Arpin, M., Algrain, M. and Louvard, D. (1994). Membrane-actin microfilament connections: an increasing diversity of players related to band 4.1. Curr. Opin. Cell Biol. 6, 136-141.[Medline]
Bennett, V. (1985). The membrane skeleton of human erythrocytes and its implications for more complex cells. Annu. Rev. Biochem. 54, 273-304.[Medline]
Birgbauer, E. and Solomon, F. (1989). A marginal band-associated protein has properties of both microtubule- and microfilament-associated proteins. J. Cell Biol. 109, 1609-1620.[Abstract]
Bossi, D. and Russo, M. (1996). Hemolytic anemias due to disorders of red cell membrane skeleton. Mol. Aspects Med. 17, 171-188.[Medline]
Bottema, C. D. and Sommer, S. S. (1993). PCR amplification of specific alleles: rapid detection of known mutations and polymorphisms. Mutat. Res. 288, 93-102.[Medline]
Chang, S. H. and Low, P. S. (2001). Regulation of the glycophorin C-protein 4.1 membrane-to-skeleton bridge and evaluation of its contribution to erythrocyte membrane stability. J. Biol. Chem. 276, 22223-22230.
Chishti, A. H., Kim, A. C., Marfatia, S. M., Lutchman, M., Hanspal, M., Jindal, H., Liu, S. C., Low, P. S., Rouleau, G. A., Mohandas, N. et al. (1998). The FERM domain: a unique module involved in the linkage of cytoplasmic proteins to the membrane. Trends Biochem. Sci. 23, 281-282.[Medline]
Cohen, W. D. (1991). The cytoskeletal system of nucleated erythrocytes. Int. Rev. Cytol. 130, 37-84.[Medline]
Cohen, W. D., Sorokina, Y. and Sanchez, I. (1998). Elliptical versus circular erythrocyte marginal bands: isolation, shape conversion, and mechanical properties. Cell Motil. Cytoskel. 40, 238-248.[Medline]
Conboy, J., Kan, Y. W., Shohet, S. B. and Mohandas, N. (1986a). Molecular cloning of protein 4.1, a major structural element of the human erythrocyte membrane skeleton. Proc. Natl. Acad. Sci. USA 83, 9512-9516.[Abstract]
Conboy, J., Mohandas, N., Tchernia, G. and Kan, Y. W. (1986b). Molecular basis of hereditary elliptocytosis due to protein 4.1 deficiency. N. Engl. J. Med. 315, 680-685.[Abstract]
Conboy, J. G. (1993). Structure, function, and molecular genetics of erythroid membrane skeletal protein 4.1 in normal and abnormal red blood cells. Semin. Hematol. 30, 58-73.[Medline]
Conboy, J. G., Chasis, J. A., Winardi, R., Tchernia, G., Kan, Y. W. and Mohandas, N. (1993). An isoform-specific mutation in the protein 4.1 gene results in hereditary elliptocytosis and complete deficiency of protein 4.1 in erythrocytes but not in nonerythroid cells. J. Clin. Invest. 91, 77-82.[Medline]
Correas, I., Leto, T. L., Speicher, D. W. and Marchesi, V. T. (1986). Identification of the functional site of erythrocyte protein 4.1 involved in spectrin-actin associations. J. Biol. Chem. 261, 3310-3315.
Culbertson, M. R. (1999). RNA surveillance. Unforeseen consequences for gene expression, inherited genetic disorders and cancer. Trends Genet. 15, 74-80.[Medline]
Davies, K. A. and Lux, S. E. (1989). Hereditary disorders of the red cell membrane skeleton. Trends Genet. 5, 222-227.[Medline]
Delaunay, J., Alloisio, N., Morle, L., Baklouti, F., Dalla, V. N., Maillet, P. and Wilmotte, R. (1996). Molecular genetics of hereditary elliptocytosis and hereditary spherocytosis. Ann. Genet. 39, 209-221.[Medline]
Detrich, H. W., III, Kieran, M. W., Chan, F. Y., Barone, L. M., Yee, K., Rundstadler, J. A., Pratt, S., Ransom, D. and Zon, L. I. (1995). Intraembryonic hematopoietic cell migration during vertebrate development. Proc. Natl. Acad. Sci. USA 92, 10713-10717.[Abstract]
Driever, W., Solnica-Krezel, L., Schier, A. F., Neuhauss, S. C., Malicki, J., Stemple, D. L., Stainier, D. Y., Zwartkruis, F., Abdelilah, S., Rangini, Z. et al. (1996). A genetic screen for mutations affecting embryogenesis in zebrafish. Development 123, 37-46.
Farr, C. J., Saiki, R. K., Erlich, H. A., McCormick, F. and Marshall, C. J. (1988). Analysis of RAS gene mutations in acute myeloid leukemia by polymerase chain reaction and oligonucleotide probes. Proc. Natl. Acad. Sci. USA 85, 1629-1633.[Abstract]
Fehon, R. G., Dawson, I. A. and Artavanis-Tsakonas, S. (1994). A Drosophila homologue of membrane-skeleton protein 4.1 is associated with septate junctions and is encoded by the coracle gene. Development 120, 545-557.
Frischmeyer, P. A. and Dietz, H. C. (1999). Nonsense-mediated mRNA decay in health and disease. Hum. Mol. Genet. 8, 1893-1900.
Gallagher, P. G., Forget, B. G. and Lux, S. E. (1998) Disorders of the erythrocyte membrane. In Nathan and Oskis Hematology of Infancy and Childhood (ed. D. G. Nathan, S. H. Orkin and F. A. Oski), pp. 544-664. Philadelphia: W. B. Saunders.
Haffter, P., Granato, M., Brand, M., Mullins, M. C., Hammerschmidt, M., Kane, D. A., Odenthal, J., van Eeden, F. J., Jiang, Y. J., Heisenberg, C. P. et al. (1996). The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio. Development 123, 1-36.
Hemming, N. J., Anstee, D. J., Staricoff, M. A., Tanner, M. J. and Mohandas, N. (1995). Identification of the membrane attachment sites for protein 4.1 in the human erythrocyte. J. Biol. Chem. 270, 5360-5366.
Hoover, K. B. and Bryant, P. J. (2000). The genetics of the protein 4.1 family: organizers of the membrane and cytoskeleton. Curr. Opin. Cell Biol. 12, 229-234.[Medline]
Knapik, E. W., Goodman, A., Ekker, M., Chevrette, M., Delgado, J., Neuhauss, S., Shimoda, N., Driever, W., Fishman, M. C. and Jacob, H. J. (1998). A microsatellite genetic linkage map for zebrafish (Danio rerio). Nat. Genet. 18, 338-343.[Medline]
Lazarides, E. (1987). From genes to structural morphogenesis: the genesis and epigenesis of a red blood cell. Cell 51, 345-356.[Medline]
Lazarides, E. and Woods, C. (1989). Biogenesis of the red blood cell membrane-skeleton and the control of erythroid morphogenesis. Annu. Rev. Cell Biol. 5, 427-452.
Leto, T. L. and Marchesi, V. T. (1984). A structural model of human erythrocyte protein 4.1. J. Biol. Chem. 259, 4603-4608.
Liao, E. C., Paw, B. H., Peters, L. L., Zapata, A., Pratt, S. J., Do, C. P., Lieschke, G. and Zon, L. I. (2000). Hereditary spherocytosis in zebrafish riesling illustrates evolution of erythroid beta-spectrin structure, and function in red cell morphogenesis and membrane stability. Development 127, 5123-5132.
Liu, S. C., Derick, L. H. and Palek, J. (1987). Visualization of the hexagonal lattice in the erythrocyte membrane skeleton. J. Cell Biol. 104, 527-536.[Abstract]
Lombardo, C. R. and Low, P. S. (1994). Calmodulin modulates protein 4.1 binding to human erythrocyte membranes. Biochim. Biophys. Acta 1196, 139-144.[Medline]
Long, Q., Meng, A., Wang, H., Jessen, J. R., Farrell, M. J. and Lin, S. (1997). GATA-1 expression pattern can be recapitulated in living transgenic zebrafish using GFP reporter gene. Development 124, 4105-4111.
Long, Q., Huang, H., Shafizadeh, E., Liu, N. and Lin, S. (2000). Stimulation of erythropoiesis by inhibiting a new hematopoietic death receptor in transgenic zebrafish. Nat. Cell Biol. 2, 549-552.[Medline]
Lorenzo, F., Dalla, V. N., Morle, L., Baklouti, F., Alloisio, N., Ducluzeau, M. T., Roda, L., Lefrancois, P. and Delaunay, J. (1994). Protein 4.1 deficiency associated with an altered binding to the spectrin-actin complex of the red cell membrane skeleton. J. Clin. Invest. 94, 1651-1656.[Medline]
Marchesi, S. L., Conboy, J., Agre, P., Letsinger, J. T., Marchesi, V. T., Speicher, D. W. and Mohandas, N. (1990). Molecular analysis of insertion/deletion mutations in protein 4.1 in elliptocytosis. I. Biochemical identification of rearrangements in the spectrin/actin binding domain and functional characterizations. J. Clin. Invest. 86, 516-523.[Medline]
Marfatia, S. M., Leu, R. A., Branton, D. and Chishti, A. H. (1995). Identification of the protein 4.1 binding interface on glycophorin C and p55, a homologue of the Drosophila discs-large tumor suppressor protein. J. Biol. Chem. 270, 715-719.
Mathias, L. A., Fisher, T. C., Zeng, L., Meiselman, H. J., Weinberg, K. I., Hiti, A. L. and Malik, P. (2000). Ineffective erythropoiesis in beta-thalassemia major is due to apoptosis at the polychromatophilic normoblast stage. Exp. Hematol. 28, 1343-1353.[Medline]
McGuire, M., Smith, B. L. and Agre, P. (1988). Distinct variants of erythrocyte protein 4.1 inherited in linkage with elliptocytosis and Rh type in three white families. Blood 72, 287-293.[Abstract]
Meng, A., Jessen, J. R. and Lin, S. (1999). Transgenesis. Methods Cell Biol. 60, 133-148.[Medline]
Mohandas, N. and Chasis, J. A. (1993). Red blood cell deformability, membrane material properties and shape: regulation by transmembrane, skeletal and cytosolic proteins and lipids. Semin. Hematol. 30, 171-192.[Medline]
Mohandas, N. and Gascard, P. (1999). What do mouse gene knockouts tell us about the structure and function of the red cell membrane? Baillieres Best. Pract. Res. Clin. Haematol. 12, 605-620.[Medline]
Mohandas, N., Phillips, W. M. and Bessis, M. (1979). Red blood cell deformability and hemolytic anemias. Semin. Hematol. 16, 95-114.[Medline]
Newton, C. R., Graham, A., Heptinstall, L. E., Powell, S. J., Summers, C., Kalsheker, N., Smith, J. C. and Markham, A. F. (1989). Analysis of any point mutation in DNA. The amplification refractory mutation system (ARMS). Nucleic Acids Res. 17, 2503-2516.[Abstract]
Nunomura, W., Takakuwa, Y., Tokimitsu, R., Krauss, S. W., Kawashima, M. and Mohandas, N. (1997). Regulation of CD44-protein 4.1 interaction by Ca2+ and calmodulin. Implications for modulation of CD44-ankyrin interaction. J. Biol. Chem. 272, 30322-30328.
Nunomura, W., Takakuwa, Y., Parra, M., Conboy, J. and Mohandas, N. (2000). Regulation of protein 4.1R, p55, and glycophorin C ternary complex in human erythrocyte membrane. J. Biol. Chem. 275, 24540-24546.
Ohanian, V., Wolfe, L. C., John, K. M., Pinder, J. C., Lux, S. E. and Gratzer, W. B. (1984). Analysis of the ternary interaction of the red cell membrane skeletal proteins spectrin, actin, and 4.1. Biochemistry 23, 4416-4420.[Medline]
Orkin, S. H. and Weiss, M. J. (1999). Apoptosis. Cutting red-cell production. Nature 401, 433-436.[Medline]
Palek, J. (1987). Hereditary elliptocytosis, spherocytosis and related disorders: consequences of a deficiency or a mutation of membrane skeletal proteins. Blood Rev. 1, 147-168.[Medline]
Palek, J. and Sahr, K. E. (1992). Mutations of the red blood cell membrane proteins: from clinical evaluation to detection of the underlying genetic defect. Blood 80, 308-330.[Medline]
Pasternack, G. R., Anderson, R. A., Leto, T. L. and Marchesi, V. T. (1985). Interactions between protein 4.1 and band 3. An alternative binding site for an element of the membrane skeleton. J. Biol. Chem. 260, 3676-3683.[Abstract]
Peters, L. L., Weier, H. U., Walensky, L. D., Snyder, S. H., Parra, M., Mohandas, N. and Conboy, J. G. (1998). Four paralogous protein 4.1 genes map to distinct chromosomes in mouse and human. Genomics 54, 348-350.[Medline]
Pootrakul, P., Sirankapracha, P., Hemsorach, S., Moungsub, W., Kumbunlue, R., Piangitjagum, A., Wasi, P., Ma, L. and Schrier, S. L. (2000). A correlation of erythrokinetics, ineffective erythropoiesis, and erythroid precursor apoptosis in thai patients with thalassemia. Blood 96, 2606-2612.
Ransom, D. G., Haffter, P., Odenthal, J., Brownlie, A., Vogelsang, E., Kelsh, R. N., Brand, M., van Eeden, F. J., Furutani-Seiki, M. et al. (1996). Characterization of zebrafish mutants with defects in embryonic hematopoiesis. Development 123, 311-319.
Ruiz-Echevarria, M. J., Gonzalez, C. I. and Peltz, S. W. (1998). Identifying the right stop: determining how the surveillance complex recognizes and degrades an aberrant mRNA. EMBO J. 17, 575-589.
Shi, Z. T., Afzal, V., Coller, B., Patel, D., Chasis, J. A., Parra, M., Lee, G., Paszty, C., Stevens, M., Walensky, L. et al. (1999). Protein 4.1R-deficient mice are viable but have erythroid membrane skeleton abnormalities. J. Clin. Invest. 103, 331-340.
Shimoda, N., Knapik, E. W., Ziniti, J., Sim, C., Yamada, E., Kaplan, S., Jackson, D., de Sauvage, F., Jacob, H. and Fishman, M. C. (1999). Zebrafish genetic map with 2000 microsatellite markers. Genomics 58, 219-232.[Medline]
Talbot, W. S. and Schier, A. F. (1999). Positional cloning of mutated zebrafish genes. Methods Cell Biol. 60, 259-286.[Medline]
Tanaka, T., Kadowaki, K., Lazarides, E. and Sobue, K. (1991). Ca2(+)-dependent regulation of the spectrin/actin interaction by calmodulin and protein 4.1. J. Biol. Chem. 266, 1134-1140.
Tchernia, G., Mohandas, N. and Shohet, S. B. (1981). Deficiency of skeletal membrane protein band 4.1 in homozygous hereditary elliptocytosis. Implications for erythrocyte membrane stability. J. Clin. Invest 68, 454-460.[Medline]
Tse, W. T. and Lux, S. E. (1999). Red blood cell membrane disorders. Br. J. Haematol. 104, 2-13.[Medline]
Wang, H., Long, Q., Marty, S. D., Sassa, S. and Lin, S. (1998). A zebrafish model for hepatoerythropoietic porphyria. Nat. Genet. 20, 239-243.[Medline]
Weed, R. I. (1970). The importance of erythrocyte deformability. Am. J. Med. 49, 147-150.[Medline]
Weinstein, B. M., Schier, A. F., Abdelilah, S., Malicki, J., Solnica-Krezel, L., Stemple, D. L., Stainier, D. Y., Zwartkruis, F., Driever, W. and Fishman, M. C. (1996). Hematopoietic mutations in the zebrafish. Development 123, 303-309.
Westerfield, M. (1993). The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Brachydanio Rerio). Eugene, OR: M. Westerfield.
Wickrema, A., Koury, S. T., Dai, C. H. and Krantz, S. B. (1994). Changes in cytoskeletal proteins and their mRNAs during maturation of human erythroid progenitor cells. J. Cell Physiol. 160, 417-426.[Medline]
Winckler, B. and Solomon, F. (1991). A role for microtubule bundles in the morphogenesis of chicken erythrocytes. Proc. Natl. Acad. Sci. USA 88, 6033-6037.[Abstract]
Woods, C. M., Boyer, B., Vogt, P. K. and Lazarides, E. (1986). Control of erythroid differentiation: asynchronous expression of the anion transporter and the peripheral components of the membrane skeleton in AEV- and S13-transformed cells. J. Cell Biol. 103, 1789-1798.[Abstract]
Woods, C. M. and Lazarides, E. (1988). The erythroid membrane skeleton: expression and assembly during erythropoiesis. Annu. Rev. Med. 39, 107-122.[Medline]
Yawata, A., Kanzaki, A., Gilsanz, F., Delaunay, J. and Yawata, Y. (1997). A markedly disrupted skeletal network with abnormally distributed intramembrane particles in complete protein 4.1-deficient red blood cells (allele 4.1 Madrid): implications regarding a critical role of protein 4.1 in maintenance of the integrity of the red blood cell membrane. Blood 90, 2471-2481.
Zhang, J., Talbot, W. S. and Schier, A. F. (1998). Positional cloning identifies zebrafish one-eyed pinhead as a permissive EGF-related ligand required during gastrulation. Cell 92, 241-251.[Medline]