3 Department of Anatomy and Cell Biology, University of Melbourne, Victoria, Australia, 3052; 4 School of Biosciences, University of Birmingham, Edgbaston, Birmingham, B15 2TT, UK; 5 Department of Cellular and Molecular Medicine, Glycobiology Research and Training Center, University of California, San Diego, La Jolla, CA 92093-0687, USA; and 6 Institute for Molecular Science of Medicine, Aichi Medical University, Japan
Received on January 1, 2002;; revised on May 2, 2002; accepted on May 29, 2002
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Abstract |
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Key words: brain development/FGF/FGF receptor/heparan sulfate/sulfotransferases
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Introduction |
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HSs are complex sulfated polysaccharides that contain polymorphic sulfated sequence motifs that are responsible for numerous protein binding and regulatory properties. HS chains are attached to core proteins to form HSPGs, which are known to have diverse biological functions (Bernfield et al., 1999), and recent genetic studies have provided compelling evidence that they are essential for normal development (Lander and Selleck, 2000
). HS chains are produced by a complex biosynthetic process that generates diverse molecular motifs with unique displays of sulfate, carboxyl, and hydroxyl groups (Lindahl et al., 1998
; Turnbull et al., 2001
). Sulfation requires a family of HS sulfotransferases (HSSTs) including four N-deacetylase-N-sulfotransferases (NDSTs; Aikawa et al., 2001
), three 6-O-sulfotransferases (6-OSTs; Habuchi et al., 2000
), at least five 3-OSTs; Shworak et al., 1999
) and a single 2-OST (Kobayashi et al., 1997
). These HSSTs have different substrate specificities and can generate different structural motifs (Aikawa et al., 2001
; Habuchi et al., 2000
; Liu et al., 1999
). Mice lacking specific HSSTs have severe phenotypes with numerous developmental defects (Bullock et al., 1998
; Forsberg et al., 1999
; Humphries et al., 1999
; Ringvall et al., 2000
). Thus, the isozyme diversity of HSSTs may be critical for generating differing sequence repertoires in HS chains.
In an earlier study we compared the structure of HS from primary cultures of neural precursor cells isolated at a mainly proliferative phase, embryonic day 10 (E10), and one where neuronal differentiation begins (E12). HS from E12 cells had longer chains with more sulfated domains, a higher level of 2-O-sulfation, and altered patterns of 6-O-sulfation and N-sulfation relative to the E10 HS (Brickman et al., 1998a,b). In this article we describe studies to determine the relative abilities of the E10 and E12 HS to activate FGF signaling through relevant receptors and to relate this to the expression levels of HSSTs in developing brain. We find that they display distinct abilities to specifically activate FGF-FGFR signaling complexes relevant to brain development, and that these changes correlate with HSST expression profiles, which vary actively both in vitro and in vivo.
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Results and discussion |
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We next determined the expression levels of the OSTs quantitatively in Nep cells using real-time PCR. Quantification of the abundance of the OST RNAs in these cells show that 2-OST RNA is the most abundant; from E10+2 to E12+2 its level increases fourfold (Figure 4A, p < 0.005). These levels are high and comparable to actin RNA levels (95% for E10+2 and 40% for E12+2 respectively; data not shown). The 6-OST RNAs also show increases in E12+2 compared to E10+2 cultures: for 6-OST1 RNA it is about twofold (Figure 4A; p < 0.005) and for 6-OST2 RNA it is about fourfold (p < 0.01). The levels of 6-OST1 RNA are higher than 6-OST2, and 6-OST3 RNA is only expressed by the E12+2 cells. This quantitative data is in good agreement with the relative intensity of the standard RT-PCR bands.
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It is interesting to compare the expression of HSSTs in the primary Nep cells with their counterparts in vivo. The OST profiles obtained from the E10+2 and E12+2 cultures are very similar to those obtained from the E10 and E12/E14 neuroepithelium, respectively (Figures 3A and 5A), as are the absolute levels of each (Figure 4A, B). Thus, there is a close correspondence between the relative expression and absolute levels of the OSTs in vivo compared with in vitro. However, the NDST expression patterns exhibit both similarities and differences. At the earlier developmental ages (compare E10+2 to E10 and E12 in vivo) the expression of NDST3 and NDST4 are similar in vivo compared with in vitro. At the later developmental ages (compare E12+2 to E12 and E14 in vivo) the expression of NDSTs 1 and 4 are similar in vivo compared with in vitro. Differences in expression are evident for NDST1 and -2 at the early stages and for NDST2 and NDST3 in the later stages. It is possible that the NDSTs are more susceptible than are OSTs to modulation of expression in response to altered cell environment.
Significance of differential HSST isozyme expression profiles in brain development
Overall these data demonstrate that the array of HSSTs expressed varies actively in developing neuroepithelial cells both in vitro and in vivo. For both the OSTs and NDSTs, the range of expressed isozymes available increases as the embryonic brain develops and correlates temporally with increasing neuronal differentiation (Caviness et al., 1995). Most important, we observed the expression of development stage-specific combinations of 6-OST and NDST isozymes.
In our earlier study of the structure of the HS from E10 and E12 neural precursor cells in vitro, we showed that the E12 HS had longer chains with a greater number of sulfated domains in comparison with the E10 HS. Although there were basic similarities in domain structure, distinct O-sulfation patterns were imposed on these domains. The difference in fine structure within the sulfated domains showed there was a higher level of 2-O sulfation and altered patterns of 6-O-sulfation and N-sulfation in the E12 HS chains in comparison to the E10 HS. Our current data is consistent with this in that we find a distinct elevation in the levels of expression of the 2-OSTs and 6-OSTs and an increasing complexity in the array of 6-OSTs expressed at the later stages. With respect to the NDSTs, there is a somewhat different trend between what we find in vivo compared with in vitro analyses. Because our earlier study analyzed HS from Nep cells in vitro, our in vitro data is the appropriate comparison here. We find increased levels of NDSTs 1 and 4 and decreasing levels of NDSTs 2 and 3 in E12 compared with E10 cultures. This is also consistent with our findings of altered patterns of N-sulfation between E10 and E12.
We suggest that these altered isozyme expression patterns would likely result in altered activity profiles of unique arrays of HSSTs; because the latter display different substrate specificities and product structures (Habuchi et al., 2000; Aikawa et al., 2001
) this could underly the observed generation of different HS structures. This leads us to propose a model in which differential expression of HSSTs results in the synthesis of variant HS species that form functional signaling complexes with FGFs and FGFRs (Figure 6). It is not yet known what specific structural characteristics of the E10 and E12 HS species are responsible for the activity differences. We speculate that variations in the repertoires of specific sequences present in the two pools of chains underlie their altered abilities to productively form ternary complexes with particular FGF-FGFR combinations. Further studies will be required to isolate and elucidate the structures of these specific functional sequences.
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These results clearly demonstrate FGF signaling specificity of two structurally distinct HS species produced by Nep cells in vitro that display different HSST expression profiles. It is plausible that the altered HSST profiles observed in vivo also result in biosynthesis of distinct HS species that could regulate activation of specific FGF-FGFR signaling complexes during brain development. Our HS preparations were derived from Nep cells in vitro, which display some differences in NDST expression compared with in vivo patterns. We do not yet know whether the latter differences are conservative with respect to synthesis of specific sequences. However, the bioactivity data we obtain with these in vitro HS species are consistent with similar potential roles for in vivo HS, and our observations align closely with the time of action of FGF2 and -8 in the developing brain.
FGF8 acts mostly prior to E12, and FGF2 acts in cortical development mainly from E10 to E14 (Fukuchi-Shimogori and Grove, 2001; Raballo et al., 2000
; see Ford-Perriss et al., 2001
). In addition, the presence of spatially specific HS species that regulate FGF-FGFR recognition has recently been described in developing mouse tissues (Allen et al., 2001
). The synthesis of these HS species may also depend on differences in the spatial localization of HSSTs. Using in situ hybridization we have evidence of differences in spatial expression of 6-OST1, 6-OST2, and 6-OST3 transcripts in distinct areas of the developing mouse brain (Drummond et al., unpublished data). Further investigation of in vivo spatiotemporal HSST expression patterns and their correlation with HS structures produced will reveal important information about the regulatory roles of HS in development.
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Materials and methods |
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RT-PCR
RT-PCR was performed on total RNA using One-Step RT-PCR (Invitrogen) according to the manufacturers instructions (1 µg total RNA mixed with 0.5 µg each of forward and reverse primers). Primers for NDST14 were as described elsewhere (Aikawa et al., 2001) and primers were designed for 2-OST and isozyme-specific primers for 6-OST13. Typical RT-PCR consisted of the following steps: 1 cycle of 94°C for 5 min and 42°C for 30 min, followed by 35 cycles of 94°C for 30 s, 50°C for 30 s, and 72°C for 45 s with a final extension of 72°C for 10 min. Primers for the 2-OST and 6-OST13 were as follows:
· 2-OST FP1: ATTAAGGAGACGGAAACAAGGAG; · 2-OST RP1: GAAGGGTGGTGACACAGTCAAG;
· 6-OST1 FP1: ACCAGCAACTCTTTCTATCCC; · 6-OST1 RP1: AGCAATACCCACCAGCATC;
· 6-OST2 FP1: TCCTTCAGACCCATTTCC; · 6-OST2 RP1: CCCACACACAGCATAACAC;
· 6-OST3 FP1: TGAATGAGAGCGAGCGGAAC; · 6-OST3 RP1: TGGATTGGAAATGAAGGCAGAG.
Each experiment was performed at least twice on at least two independent RNA preparations, and figures show representative examples.
Real-time quantitative PCR of OSTs
The reverse transcription step was carried out according to manufacturers instructions using the SuperscriptIITM Preamplification system (Invitrogen) except that 50 ng of total RNA was used in a 10-µl total reaction volume. Triplicate 2-µl samples of this cDNA were quantified by real-time PCR using a Corbett RG-2000 (Corbett, Sydney, Australia) using the parameters detailed in Table I. Three to four independent dissections or cell culture experiments were analyzed, and triplicate real-time measurements were performed for each cDNA preparation. Data were analyzed using the RG-2000 quantification and melt analysis programs. Reactions were performed in a 20 µl volume: 2 µl of cDNA (from a typical RT-PCR reaction as described) and 18 µl of a master mix containing 0.2 U Taq polymerase (Fisher Biotech), buffer, 0.5 µM forward and reverse primers, MgCl2 optimized at either 2 mM or 3 mM (see Table I) and 0.5x SYBR Green (Molecular Probes). A typical protocol was 1 min at 94°C (one cycle), 15 s at 94°C, 20 s at 55°C, 30 s (2-OST, 6-OST3) and 20 s (6-OST1, 6-OST2) at 72°C, 10 s at 87°C (for detection of the fluorescent product) for 35 cycles. Some additional primers were used and compared with those used for standard RT-PCR. These were 6-OST1 FP2: CTGCATCTTCTTACCCTTTAC; 6-OST2 FP3: GGTCAGAATCTGAGTCAGAATC; 6-OST3 RP2: CCAAAGTAATCCAAGAGAAG.
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Semi-quantitative RT-PCR of FGFR b and c splice variant expression
The methodology for both cDNA synthesis and subsequent PCR were essentially as already described. The FGFR splice variant RT-PCR strategy used primers as follows.
Forward for loop III PCR: · FGFR1b, CTTGACGTCGTGGAACGATCT;
· FGFR1c, CTTGACGTCGTGGAACGATCT;
· FGFR2b, CCCATCCTCCAAGCTGGACTGCCT;
· FGFR2c, CCCATCCTCCAAGCTGGACTGCCT; · FGFR3b GACATACACACTGGATGTGCTGGA;
· FGFR3c, GACATACACACTGGATGTGCTGGA; · FGFR4, CAACTCCATCGGCCTTTCCTACCA.
Reverse (for Loop III cDNA synthesis and PCR):
· FGFR1b, CTGGTTAGCTTCACCAATAT; · FGFR1c, TTCCAGAACGGTCAACCATGCAGA;
· FGFR2b, ATCTGGGGAAGCCGTGATCTCCTT; · FGFR2c, TGGCAGAACTGTCAACCATGCAGA;
· FGFR3b, GGCCTTCTCAGCCACGCCTAT; · FGFR3c, AGCACCACCAGCCACGCAGAGTGA;
· FGFR4, GGCAGGTCTAGATTCACAAGGCCC.
Internal (5', for exon-specific PCR): · FGFR1b, CGGGAATTAATAGCTCGGAT;
· FGFR1c, ACTGCTGGAGTTAATACCACCGAC; · FGFR2b, CTGAAGCACTCGGGGATAAATAGC;
· FGFR2c, GGTGTTAACACCACGGACAAAGAG; · FGFR3b, GAATGTGGAGGCAGACGCACG;
· FGFR3c, TGCAGGCGCTAACACCACCGACAA; · FGFR4, CAGGCTCACTGGTTCTGCTTGTGC.
The design allows for RT-PCR expression analysis of the b or c splice variant using the Loop III forward and reverse primers and for subsequent Southern analysis of the PCR products for verification of identity using the internal 5' b or c exon-specific primers as probes. PCR products were transferred to a Hybond N+ (Amersham) membrane and hybridized at 42°C overnight with their corresponding internal primers that had been end-labeled with -32P-ATP and polynucleotide kinase (Southern conditions and end-labeling detailed in Ford et al., 1997
). To quantify the expression levels of each gene the intensities of the PCR product bands at 30 cycles were measured by exposing the blots to a PhosphorImager screen (Fujix Imaging Plate, Type BAS-IIIS) and using MacBasII software. Analyses were performed in duplicate on duplicate RNA preparations from the different ages and signals quantified using a Fujix PhosphorImager and MacBasII software. The results represent mean values ± SD.
HS purification and BaF3 assays
HS was purified from primary cultures of Nep cells as described (Brickman et al., 1998a,b). BaF3 lymphoid cells expressing various FGFR isoforms (Ornitz et al., 1996
) were incubated for 72 h with 1 nM FGF1, FGF2, or FGF8 (b isoform) and HS from E10+2 or E12+2 cells. Cell numbers were measured using an MTT (3-[4,5-Dimethylthiazol-2-yl]-2,5 diphenyltetrazolium bromide) assay as described (Guimond and Turnbull, 1999
). Data is presented as mean ± SD.
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Acknowledgments |
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Abbreviations |
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Footnotes |
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2 To whom correspondence should be addressed; E-mail: j.e.turnbull@bham.ac.uk
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References |
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