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Distinct transcriptional regulatory mechanisms underlie left-right asymmetric expression of lefty-1 and lefty-2.

Both lefty-1 and lefty-2 genes are expressed on the left side of developing mouse embryos and are implicated in left-right (L-R) axis formation. With the use of transgenic analysis, the transcriptional regulatory regions of these genes responsible for their L-R asymmetric expression have now been investigated. The 9.5-kb upstream region of lefty-1 and the 5.5-kb upstream region of lefty-2 reproduced the expression pattern of the corresponding gene. Examination of deletion constructs revealed the presence of a left side-specific enhancer (ASE) that is essential and sufficient for lefty-2 asymmetric expression. In contrast, the asymmetric expression of lefty-1 was shown to be determined by a combination of bilateral enhancers and a right side-specific silencer (RSS). The 9. 5-kb region of lefty-1 and the 5.5-kb region of lefty-2 responded to iv and inv, upstream genes of lefty-1 and lefty-2. The regulation of lefty-2 by iv and inv was mediated by ASE. These results suggest that, in spite of the similarities between lefty-1 and lefty-2, different regulatory mechanisms underlie their asymmetric expression.

Vertebrates exhibit numerous left-right (L-R) asymmetries, such as the positions of the heart and spleen on the left side of the body. In rodents, L-R asymmetry appears to be determined at the presomite stage (Brown et al. 1991;Fujinaga and Baden 1991a,b). One of the earliest morphological indicators of laterality is the looping of the cardiac tube toward the right side. This event is accompanied by a clockwise axial rotation referred to as embryonic turning, and is followed, much later, by the asymmetric orientation of various visceral organs.
The molecular mechanism that underlies L-R axis formation has remained largely unknown until recently, when insights into this important process have been gained (for review, see King and Brown 1997;Varlet and Robertson 1997;Levin and Mercola 1998). The entire process by which L-R asymmetry is established can be divided into three phases: (1) the initial determination of L-R polarity, (2) L-R asymmetric expression of signaling molecules, and (3) L-R asymmetric morphogenesis induced by these signaling molecules. The mechanism of the initial determination of L-R polarity remains unknown, although several models have been proposed (Brown and Wolpert 1990;Yost 1992). In mice, several mutations are known to affect L-R patterning (Hummel and Chapman 1959;Yokoyama et al. 1991;Heymer et al. 1997;Melloy et al. 1997), two of the more extensively studied of which are iv and inv. L-R specification is randomized in iv/iv mice, whereas it is reversed in inv/inv mice. These phenotypes suggest that the iv and inv genes likely contribute to the initial (early) determination of L-R polarity. The iv and inv genes have been shown to encode a protein related to axonemal dynein (Supp et al. 1997), and a novel protein with ankyrin repeats (Mochizuki et al. 1998), respectively. However, the precise function of these proteins remains unclear.
With regard to the second phase of L-R axis formation, the L-R asymmetric expression of signaling molecules, Levin et al. (1995) have shown that the activin receptor typeIIA, sonic hedgehog, and cNR-1 genes are expressed asymmetrically in chick embryos, and they have proposed a hierarchy for these genes. cNR-1 is a chick homolog of mouse nodal (Levin et al. 1995), which belongs to the TGF␤ superfamily. The pattern of nodal expression is conserved among vertebrates; it is expressed in the left half of mouse, chick and Xenopus embryos (Collignon et al. 1996;Lowe et al. 1996). We have identified previously two highly related TGF␤-related genes, lefty-1 and lefty-2, that are also expressed on the left side of developing mouse embryos (Meno et al. 1996(Meno et al. , 1997. Whereas lefty-1 is expressed predominantly in the prospective floor plate (PFP), lefty-2 is expressed in lateral plate mesoderm (LPM). Misexpression experiments in chick and Xenopus, as well as analyses of knockout mice support the notion that these genes encode signaling molecules for L-R positional information (Levin et al. 1997;Sampath et al. 1997;Meno et al. 1998). For example, mice deficient in lefty-1 show a variety of positional defects in visceral organs, indicating that lefty-1 is required for the correct positioning of these organs ). In lefty-1 −/− mice, lefty-2 and nodal are ectopically expressed on the right side, suggesting that Lefty-1 restricts the expression of nodal and lefty-2 to the left side, perhaps by acting as (or inducing) a midline barrier. The observation that expression of lefty-1, lefty-2, and nodal is affected in iv and inv mutant embryos suggests that lefty-1, lefty-2, and nodal all act downstream of the iv and inv genes.
With regard to the third phase of establishment of L-R asymmetry, situs-specific morphogenesis in response to L-R asymmetric signals, genetic evidence in mice (Oh and Li 1997) suggests that type II activin receptors (ActRIIA and ActRIIB) serve as functional receptors for Nodal (and also for Lefty-2). Furthermore, recent studies (Logan et al. 1998;Meno et al. 1998;Piedra et al. 1998;Ryan et al. 1998;Yoshioka et al. 1998) have indicated a role for Pitx2 (also known as Rieg, Otlx2, and Brx1;Mucchielli et al. 1996;Semina et al. 1996;Gage and Camper 1997;Kitamura et al. 1997), a bicoid-type homeobox transcription factor, in mediating transmission of signals from Nodal and Lefty-2. In addition, Snr-1, which encodes a zinc finger-type transcription factor, is expressed on the right side of chick embryos (Isaac et al. 1997) and may be negatively regulated by Nodal.
In spite of the recent progress, our knowledge of L-R determination remains limited. For example, the mechanism by which the initial determination of L-R polarity eventually results in the asymmetric expression of signaling molecules is not known. An understanding of the early events that take place before the asymmetric expression of lefty-1, lefty-2, and nodal will require elucidation of the mechanisms by which these genes are regulated. We have now identified the transcriptional regulatory elements of lefty-1 and lefty-2 that are responsible for the asymmetric expression of these genes. Our results indicate that lefty-2 is regulated by an asymmetric enhancer that is active exclusively on the left side. In contrast, lefty-1 is controlled by an asymmetric silencer that represses expression on the right side. These observations suggest that, although both lefty-1 and lefty-2 are expressed asymmetrically, different regulatory mechanisms underlies the L-R asymmetric transcription of the two genes.

Results
The upstream regions of lefty-1 and lefty-2 determine the expression patterns of these genes lefty-1 and lefty-2 are tightly linked on mouse chromosome 1, being separated by a distance of only 30 kb ( Fig.   1A; Meno et al. 1997). Both genes exhibit similar exonintron structures and possess a putative TATA box at an expected location. We first examined the transcriptional activity of upstream regions of lefty-1 and lefty-2. For lefty-1, the 9.5-kb upstream region containing the TATA box (from −9.5 kb to +80 bp relative to the TATA box) was linked to lacZ, yielding L1-9.5 (Fig. 1A). For lefty-2, the 5.5-kb upstream region including the TATA box (−5.5 kb to +90 bp relative to the TATA box) was fused to lacZ, yielding L2-5.5 (Fig. 1A). L1-9.5 and L2-5.5 were injected separately into the pronuclei of fertilized embryos at the one-cell stage. The embryos were allowed to develop in utero until embryonic day 8.2 (E8.2), at which time they were recovered and the expression of the lacZ transgene was examined by staining with X-gal (5bromo-4-chloro-3-indoyl-␤-D-galactoside). Embryos injected with L1-9.5 showed marked X-Gal staining in the PFP (Fig. 1B,C). Transverse sections of the embryos indicated that X-gal staining was localized predominantly in the left half of the PFP (Fig. 1D,E). Less marked X-gal staining was also detected in the anterior region of the LPM on the left side ( Fig. 1B-E) This pattern of X-gal staining was indistinguishable from that of the distribution of lefty-1 mRNA determined previously by whole mount in situ hybridization (Meno et al. 1997).
Embryos injected with L2-5.5 showed marked X-gal staining in left LPM as well as weaker X-gal staining in left PFP (Fig. 1F-I). At E7.0, X-gal staining was detected in the emerging mesoderm (data not shown). These staining patterns were identical to the distribution profiles of lefty-2 mRNA determined previously by whole mount in situ hybridization (Meno et al. 1997). Thus, L1-9.5 and L2-5.5 were able to recapitulate the expression patterns of lefty-1 and lefty-2, respectively. These results also indicate that lefty-1 and lefty-2 are regulated independently, even though they are tightly linked on the same chromosome.

L-R asymmetric expression of lefty-2 is controlled by a left side-specific enhancer
To locate cis elements responsible for the asymmetric expression of lefty-2, we tested various restriction fragments derived from the 5.5-kb upstream region for the ability to confer asymmetric expression on lacZ ( Fig.  2A). Whereas the 5.5-kb upstream fragment fully retained this ability, asymmetric expression was not apparent with L2-3.3 or L2-1.6 ( Fig. 2A), suggesting the presence of a left side-specific enhancer in the 2.2-kb region between −5.5 and −3.3 kb. Although L2-3.3 embryos failed to show X-gal staining in LPM, symmetrical staining was detected in the paraxial mesoderm. Because X-gal staining in the paraxial mesoderm was not apparent in L2-1.6 embryos ( Fig. 2A), a paraxial mesodermspecific enhancer (PME; Fig. 2) is likely located between −3.3 and −1.6 kb.
To delineate further the cis elements responsible for asymmetric expression, we prepared two sets of deletion mutants from L2-5.5, all of which contained the 3.3-kb proximal upstream region (Fig. 2B). Among the 5Ј-dele-tion mutants, left-sided lacZ expression in LPM and PFP was observed with L2-5Ј⌬25 and L2-5Ј⌬75 but not with L2-5Ј⌬199 (Fig. 2B,C). Of the 3Ј-deletion mutants, L2-3Ј⌬56 showed asymmetric expression, whereas L2-3Ј⌬37 did not (Fig. 2B,C). These results indicated the presence of a left side-specific enhancer referred to as ASE (asymmetric enhancer), in a 380-bp region between −4.1 and −3.7 kb. The observation that X-gal staining in the paraxial mesoderm was often apparent when ASE was deleted (L2-5Ј⌬199 and L2-3Ј⌬37; Fig. 2B) suggest that ASE may be required not only for left-sided expression but also for suppressing the paraxial mesoderm-specific enhancer.
Internal deletion of the 380-bp region containing ASE from L2-5.5, yielding L2⌬37-199 (Fig. 3A), abolished asymmetric expression (Fig. 3D,G), indicating that ASE is absolutely required for such expression. The ASE-lef2p construct (Fig. 3A), in which the 380-bp ASE fragment is linked to the minimal promoter region of lefty-2 (from −300 to +90 bp), yielded left-sided X-gal staining in left LPM and PFP (Fig. 3B,E). Finally, constructs in which the 380-bp fragment was linked to the hsp68 promoter instead of the lefty-2 promoter [ASE-hsp (R) and ASEhsp (S); Fig. 3A] gave rise to X-gal staining in left LPM and left PFP (Fig. 3C,F). These results demonstrate that ASE is essential and sufficient for the asymmetric expression of lefty-2.

The left side-specific enhancer of lefty-2 is composed of multiple subdomains
The 380-bp region containing ASE was further dissected by directional and internal deletion analysis (Fig. 4A,B). Serial deletion of the 380-bp region from the 5Ј end resulted in the sequential loss of lacZ expression in the PFP and LPM; expression disappeared first from the PFP, then from the posterior LPM, and finally from the anterior LPM (Fig. 4A,C). The L2-5Ј⌬501 construct was expressed in LPM but showed virtually no expression in the PFP. The mutants L2-5Ј⌬98, L2-5Ј⌬505, and L2-5Ј⌬502 showed asymmetric expression in the anterior portion of LPM, but they exhibited a gradual loss of expression in the posterior LPM as the extent of the de- letion increased; the graded pattern of X-gal staining along the anteroposterior (A-P) axis was most evident with L2-5Ј⌬505 and L2-5Ј⌬502 (Fig. 4C). Finally, with L2-5Ј⌬503 and L2-5Ј⌬504, expression in the anterior LPM was virtually abolished (Fig. 4A).
Serial deletion of the 380-bp region from the 3Ј end resulted in the sequential loss of expression in the anterior LPM (L2-3Ј⌬12 and L2-3Ј⌬54) followed by that in the posterior LPM and PFP (L2-3Ј⌬52). These results suggest that ASE is composed of multiple (at least three) subdomains; in the 5Ј to the 3Ј direction, a subdomain that controls expression in the PFP, one for the posterior portion of LPM, and one for the anterior portion of LPM. Consistent with the results obtained with the directional deletion mutants, internal deletion mutants lacking the 3Ј portion of the 380-bp region (L2-⌬54-504 and L2- ⌬54-199) showed X-gal staining in the posterior portion of LPM but lost that in the anterior portion of LPM (Fig. 4C).

Regulation of lefty-1 by bilateral enhancers and a right side-specific silencer
To localize the transcriptional regulatory elements of lefty-1, we tested various restriction fragments derived from the 9.5-kb upstream region for the ability to confer asymmetric expression (Fig. 5A). The 6.0-kb upstream fragment (L1-6.0) fully retained the ability to mediate lacZ expression in the PFP and left LPM (Fig. 5B,E). Whereas L1-3.0 and L1-1.3 retained asymmetric transcriptional activity in the PFP [transverse section indicated that X-gal staining in PFP was left sided (data not shown)], no expression was apparent in LPM ( Fig.  5C,D,F,G). These results suggested the presence of an LPM-specific enhancer (LPE) in the 3.0-kb region between −6.0 and −3.0 kb.
The asymmetric expression in the PFP was maintained in L1-3.0 and L1-1.3 (Fig. 5), suggesting the presence of a PFP-specific enhancer(s) within the 1.3-kb upstream region. To delineate further the position of the PFP-specific enhancer, we constructed a series of 5Ј-deletion mutants from L1-1.3. Because the asymmetric ex-  (PME) paraxial mesoderm-specific enhancer. (C) Anterior views (top) and lateral views (bottom) of X-gal stained embryos transgenic for the indicated constructs. Marked X-gal staining in left LPM is apparent with 5Ј⌬75 and 3Ј⌬56, but such staining was not detected with 5Ј⌬199 or 3Ј⌬37. Both 5Ј⌬199 and 3Ј⌬37 show weak bilateral X-gal staining in the paraxial mesoderm, which is probably due to the presence of PME in the proximal upstream region.
pression of lacZ in the PFP was not apparent with L1-5Ј⌬9 or L1-5Ј⌬8, the PFP-specific enhancer referred to as NPE (neural plate enhancer) was localized to a 200-bp region between −1.3 and −1.1 kb.
To examine whether LPE and NPE possess L-R specificity, LPE (the 1.5-kb fragment from −4.5 to −3.0 kb) and NPE (the 0.2-kb fragment from −1.3 to −1.1 kb) were linked to the hsp68 promoter, yielding LPE-hsp and NPE-hsp, respectively (Fig. 6A). Unexpectedly, LPE-hsp gave rise to X-gal staining on both sides of LPM (Fig. 6B). As with L1-9.5 (Fig. 1D,E), the anterior portion, but not the posterior portion, of LPM was positive for X-gal staining. NPE-hsp also lacked L-R specificity, with bilateral X-gal staining apparent in a broad region of the neural plate along the A-P axis (Fig. 6C,F,I,M). Therefore, unlike ASE of lefty-2, neither LPE nor NPE alone confers L-R specificity. It was most likely that LPE and NPE are bilateral enhancers, and that L-R specificity was determined by some other cis element.
Because L1-1.3 showed left-sided X-gal staining in the PFP (Fig. 5A,D,G), a cis element that controls asymmetric expression appeared to reside in the proximal promoter region (between −1.3 and +80 bp). When LPE was linked to an 0.9-kb proximal upstream region (−0.9 kb to +80 bp) of lefty-1 (LPE⌬8), left-sided X-gal staining was apparent in the anterior portion of LPM (Fig. 6B). However, when the proximal promoter region was further deleted toward the TATA box (LPE⌬10 containing LPE and an upstream fragment from −70 to +80 bp), X-gal staining in LPM was bilateral (Fig. 6B). These results suggested the presence of a right side-specific silencer referred to as RSS (right side-specific silencer), in the proximal promoter region (between −0.9 kb and −70 bp).
Similar experiments were performed with a set of constructs containing NPE. In contrast to the L-R specificity of L1-1.3 (Fig. 5), the deletion of the proximal promoter region between −0.9 kb and −70 bp (NPE⌬10) resulted in bilateral lacZ expression throughout the neural plate (Fig. 6A,D,G,J,N), with the exception that left-sided X-gal staining was maintained in the most anterior region. Furthermore, the addition of the 0.8-kb region (from −0.9 kb to −70 bp) to NPE-hsp, yielding NPE-RSShsp, rescued left-sided expression in the PFP (Fig.  6E,H,K,O); when compared with NPE-hsp, NPE-RSShsp gave rise to left-sided X-gal staining in the PFP with the exception of the middle portion of the PFP. These results again suggested the presence of an RSS element in the 0.8-kb region between −0.9 kb and −70 bp. We therefore concluded that the asymmetric expression of lefty-1 in the PFP and LPM is determined not by enhancers but by RSS, a silencer that can repress transcription on the right side when combined with the bilateral enhancers.
Transcriptional regulatory elements of lefty-1 and lefty-2 respond to iv and inv mutations Expression of lefty-1 and lefty-2 is affected in situs mutants such as iv/iv and inv/inv mice; the pattern of expression is randomized (left-sided, right-sided, or bilateral) in iv/iv mutants, and reversed in inv/inv mutants (Collignon et al. 1996;Lowe et al. 1996;Meno et al. 1996Meno et al. , 1997. Therefore, lefty-1 and lefty-2 both act downstream of the iv and inv genes. We examined whether expression of the lefty-1-lacZ and lefty-2-lacZ transgenes was affected by the iv and inv mutations, by establishing permanent transgenic lines containing L1-9.5 or L2-5.5 (lines 1-1 and 2-1, respectively). Embryos derived from these permanent lines showed X-gal staining patterns identical to those observed in the transient transgenic embryos; line 1-1 embryos showed marked X-gal staining in left PFP as well as weaker staining in left LPM, whereas line 2-1 embryos exhibited marked X-gal staining in left LPM and weaker staining in left PFP (data not shown).
Line 2-1 mice were mated with iv/iv mice and the lacZ transgene was transferred to the iv/iv background. The iv/iv embryos harboring the L2-5.5 transgene showed three different patterns of X-gal staining in LPM; left-sided, right-sided, or bilateral ( Fig. 7A-C). The L2-5.5 transgene was also transferred to the inv/inv background by mating line 2-1 animals with inv/+ mice (inv/inv homozygotes are lethal). Among the embryos obtained by crossing inv/+, L2-5.5 mice with inv/+ mice, ∼25 % of lacZ + embryos (14/46), which were inv/inv homozygotes, showed abberant X-gal staining pattern in LPM; the staining pattern in such embryos was either rightsided (11/14 embryos; Fig. 7F) or bilateral (3/14 embryos; Fig. 7E). These X-gal staining patterns observed in the iv/iv and inv/inv mice were identical to the patterns of lefty-2 mRNA distribution in these mutants (Meno et al. 1997). Therefore, we concluded that the 5.5-kb upstream region of lefty-2 can reproduce the response to the iv and inv mutations.
To examine whether ASE alone can respond to the iv mutation, we also established a permanent line containing ASE-hsp (line 2-2). Line 2-2 embryos expressed lacZ in left LPM and PFP (data not shown), as observed with the transient transgenic embryos (Fig. 3C,F). Line 2-2 mice were then crossed with iv/iv mice and inv/+ mice. The iv/iv embryos harboring an ASE-hsp transgene showed either left-sided, bilateral, or right-sided X-gal staining both in LPM and PFP (Fig. 7G-I). The inv/inv embryos harboring the ASE-hsp transgene showed either right-sided or bilateral X-gal staining in LPM and PFP (Fig. 7K,L). These results indicate that the regulation of lefty-2 by iv and inv genes is mediated through ASE. Similarly, the L1-9.5 transgene was transferred to the inv/inv and iv/iv backgrounds. The iv/iv embryos containing the L1-9.5 transgene showed either left-sided (Fig. 7M,P), right-sided (Fig. 7O,R), or bilateral (Fig.  7N,Q) expression in the PFP and LPM. When inv/+, L1-9.5 mice were mated with inv/+ mice, most of the transgene-positive inv/inv embryos showed right-sided X-gal staining in the PFP (Fig. 7U,X), whereas the remainders exhibited bilateral expression in the PFP (Fig. 7T,W). These results indicate that the regulation of lefty-1 by iv and inv genes is mediated by the 9.5-kb upstream region.

Discussion
Regulation of lefty-1 and lefty-2 by distinct mechanisms lefty-1 and lefty-2 are tightly linked on mouse chromosome 1. However, we have now demonstrated that the two genes are regulated independently and by distinct mechanisms (Fig. 8). The asymmetric expression of lefty-2 appears to be determined solely by a left sidespecific enhancer (ASE), given that ASE alone is necessary and sufficient for L-R asymmetric expression. In contrast, L-R asymmetric expression of lefty-1 is determined not by the enhancers (LPE and NPE) but by an RSS.
Despite the similarities in structure and expression patterns between lefty-1 and lefty-2, several differences are apparent between the two genes. First, their major expression domains differ; whereas lefty-1 is expressed predominantly in the PFP, lefty-2 is expressed predominantly in LPM. In addition, lefty-1 expression slightly precedes that of lefty-2; lefty-1 expression begins in the PFP at the two-somite pair stage, whereas the asymmetric expression of lefty-2 begins in left LPM at the threesomite pair stage (Meno et al. 1997;Heymer et al. 1997). Our data suggested that, although both are expressed in

GENES & DEVELOPMENT 265
Cold Spring Harbor Laboratory Press on July 22, 2018 -Published by genesdev.cshlp.org Downloaded from left LPM, lefty-1 and lefty-2 are regulated differently. This difference may reflect distinct cell lineages for lefty-1-expressing and lefty-2-expressing mesoderm cells; lefty-2 expression in LPM begins at the left side of the node, whereas lefty-1 expression in LPM begins at the left foregut. Expression of lefty-1 and that of lefty-2 are affected similarly in iv and inv mutants (Meno et al. 1997) but differently in other mutants such as nt and ft. In nt mutant mice, for instance, lefty-1 is not expressed, whereas lefty-2 expression is bilateral (Melloy et al. 1998). A second difference between lefty-1 and lefty-2 is that they likely play distinct roles in L-R determination. Recent analysis of lefty-1 −/− mutant mice has suggested that the role of lefty-1 is to restrict the expression of lefty-2 and nodal to the left side of the embryos, whereas lefty-2 and nodal may encode signals for leftness ). Thus, lefty-1 appears to act upstream of lefty-2 and nodal. These differences between the two genes may explain, at least in part, the existence of distinct mechanisms for the regulation of lefty-1 and lefty-2.
The expression patterns of nodal and lefty-2 are highly similar; the timing and sites of their expression are in-distinguishable. Furthermore, these two genes may play similar roles in L-R determination; as mentioned above, both Lefty-2 and Nodal may act as determinants for leftness ). In addition, both Lefty-2 and Nodal can induce Pitx2 expression in chick embryos (Logan et al. 1998;Piedra et al. 1998;Yoshioka et al. 1998). Therefore, the asymmetric expression of nodal and lefty-2 may be regulated by similar mechanisms. We and others have recently identified a left side-specific enhancer in the nodal gene (H. Adachi, Y. Saijoh, S. Ohishi, K. Mochida, and H. Hamada unpubl.; E.J. Robertson, pers. comm.).
Our data suggest that ASE is composed of multiple subdomains. Thus, it is likely that different sets of cisregulatory sequences within the 380-bp region contribute to the regulation of lefty-2 in the PFP, the anterior LPM, and the posterior LPM. This notion is supported by our observations with lefty-1 mutant mice ). In lefty-1 −/− embryos, both lefty-2 and nodal are bilaterally expressed in LPM. However, the ectopic expression of lefty-2 in right LPM was always confined to the anterior portion, suggesting that lefty-2 is regulated differently in the anterior and posterior LPM. Regulation of lefty-1 and lefty-2 by signals downstream of iv and inv Given that the expression of lefty-1, lefty-2, and nodal is affected in iv/iv and inv/inv mutant embryos (Collignon et al. 1996;Lowe et al. 1996;Meno et al. 1996Meno et al. , 1997, these three genes all likely act downstream of the iv and inv genes. Thus, signals downstream of the iv and inv genes must reach the transcriptional regulatory regions of lefty-1 and lefty-2. The 5.5-kb upstream region of lefty-2 was able to respond to the iv and inv mutations. Furthermore, ASE-hsp lacZ also responded to these mutations. Therefore, signals downstream from the iv and inv genes likely target the 380-bp region of ASE. The 9.5-kb upstream region of lefty-1 was also able to respond to the iv and inv mutations. Regulation of lefty genes by the iv and inv genes must be indirect because these latter two genes appears to act at an earlier stage of L-R determination. Furthermore, the iv gene encodes a protein related to axonemal dynein, not a transcription factor (Supp et al. 1997). Clarification of the mechanism by which signals downstream of iv and inv regulate lefty genes will require identification of transcription factors that interact with ASE of lefty-2 and those that bind to RSS of lefty-1. The 380-bp region of ASE and the 0.7-kb region of RSS each possess a number of potential binding sequences for known transcription factors. To identify transcription factors that bind to ASE and RSS, detailed mutational analysis of ASE and RSS as well as the sequence information on other asymmetric enhancers (such as the ones found in nodal gene and human lefty-2) are needed.

Positive signals on the left side or negative signals on the right side?
Theoretically, L-R asymmetric transcription of left sideexpressed genes can be achieved by one of the two regulatory mechanisms; transcriptional activation on the left side or transcriptional silencing on the right side. Although our data may appear to support the former mechanism for lefty-2 and the latter mechanism for lefty-1, it is premature to reach such a conclusion. In the case of lefty-2, for instance, the L-R specificity of ASE could be conferred either by the presence of an ASEactivating signal on the left side and its absence on the right side, or by the presence of an ASE-repressing signal on the right and its absence on the left. The same logic applies to RSS of lefty-1. Distinguishing between the two possible mechanisms for the asymmetric expression of lefty-1 and lefty-2 will require identification of the transcription factors that bind to ASE and RSS.

Materials and methods lacZ constructs
Chromosomal lefty-1 and lefty-2 genes were isolated from a genomic library derived from 129/Sv mice. The 3Ј end of the lefty-1-upstream fragment (SalI-HindIII 9.5 kb) is located in the 5Ј-untranslated region (+80 bp relative to the putative TATA box, −20 bp relative to the translation initiation codon). The 3Ј end of the lefty-2 upstream fragment (SalI-HindIII 6 kb) is also located in the 5Ј-untranslated region (+90 bp relative to the putative TATA box, −10 bp relative to the initiation codon). These upstream fragments were linked to the lacZ fragment (HindIII-BamHI 4 kb) derived from pCH110 (Pharmacia) and were then subcloned in Bluescript or pGEM, yielding L1-9.5 and L2-5.5. In the 5Ј⌬ set of L2-5.5 deletion mutants, deletion proceeded from the SalI site located at the 5Ј end of L2-5.5 toward the 3Ј end. In the 3Ј⌬ set of L2-5.5 deletion mutants, deletion proceeded from the internal BamHI site (located at −3.3 kb) toward the 5Ј end (3Ј-deletion mutants were constructed by adding each of 3Ј-deleted fragments to L2-3.3 in a correct orientation). For generation of lacZ constructs containing the hsp68 promoter, various DNA fragments were rendered blunt ended and subcloned at the SmaI site of hsp68 lacZpA [Kothary et al. 1989;Logan et al. 1998; a generous gift from A. Joyner (New York University Medical Center) and J. Rossant (Samuel Lunenfeld Research Institute, Mount Sinai Hospital)]. For all the constructs, lacZ fragments free of vector sequences were isolated by gel electrophoresis and used for microinjection.

Transient transgenic assay
Transgenic mice were generated by pronuclear injection of lacZ fragments (4 ng/µl) into fertilized eggs obtained from intercrosses between (C57BL/6× C3H)F1 mice (Hogan et al. 1994;Sasaki and Hogan 1994). The injected embryos were transferred into pseudopregnant recipients, and allowed to develop in utero until E8.2. Because of the narrow window of lefty expression, it was necessary to recover the embryos at a stage between three and eight somite pairs, for which the recipient mice were anesthetized with Nenbutal and the stage of embryos was estimated by a size of the decudia. The E8.2 embryos were examined for the presence of the transgene (by PCR) and for lacZ expression Cold Spring Harbor Laboratory Press on July 22, 2018 -Published by genesdev.cshlp.org Downloaded from (by X-gal staining). The activity of ␤-galactosidase in dissected embryos was detected by a standard protocol (Hogan et al. 1994). For each construct, >120 embryos were microinjected and transferred, and ജ10 transgene-positive embryos were examined by X-gal staining. The amount of ␤-galactosidase activity was estimated from the extent of X-gal staining after incubating in the staining buffer for 2, 8, and 24 hr. Primers used to amplify the lacZ sequence were 5Ј-TTGCCGTCTGAATTTGACCTG-3Ј and 5Ј TCTGCTTCAATCAGCGTGCC-3Ј (Sasaki and Hogan 1996).

Permanent transgenic lines
For some lacZ fragments, the transgenic embryos were allowed to develop to term and permanent transgenic lines were established. The presence of the transgene was examined by Southern blot analysis with the lacZ fragment as probe. The lacZ transgenes were introduced into the background of iv/iv or inv/ +, by first mating transgenic mice with iv/iv and inv/+ mice, respectively. The genotype of the resulting offsprings was determined by PCR or Southern blotting analysis. iv/iv, lacZ or inv/+, lacZ mice thus obtained were mated, and the expression of the transgenes at E 8.2 was examined. The iv alleles were genotyped by PCR with the primers, 5Ј-GCCAGCAATGAAC-GAGTGGCCCTCAAACCT-3Ј and 5ЈAGCTCTGGAAACC-GTGGCTGGTGTGGCTGT-3Ј, followed by TaqI digestion. The wild-type iv allele is sensitive to TaqI, yielding 50-bp fragments, whereas the mutant allele is resistant to TaqI, yielding a 100-bp fragment (Supp et al. 1997). The inv alleles were also genotyped by PCR. A pair of primers, 5Ј-GTACACACCCCTT-GATTATGCA-3Ј and 5Ј-TTGGCTGCAGCGTCTTTCCTCA-3Ј were used to detect the wild-type inv allele. These primers would amplify a 212-bp fragment that is located in exon 11 and is deleted in the mutant allele. The primers used to detect the mutant inv allele are 5Ј-TCAGAAGAGTATAATAGCCATC-3Ј and 5Ј-TGTCCACAAAAGCATGGTGAAGAAG-3Ј. These primers would amplify a 348-bp fragment derived from the tyrosinase transgene.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Chemistry

text:
Phosphorylation of Heterochromatin Protein 1 by Casein Kinase II Is Required for Efficient Heterochromatin Binding inDrosophila *

Heterochromatin-associated protein 1 (HP1) is a nonhistone chromosomal protein with a dose-dependent effect on heterochromatin mediated position-effect silencing. It is multiply phosphorylated in vivo. Hyperphosphorylation of HP1 is correlated with heterochromatin assembly. We report here that HP1 is phosphorylated by casein kinase II in vivo at three serine residues located at the N and C termini of the protein. Alanine substitution mutations in the casein kinase II target phosphorylation sites dramatically reduce the heterochromatin binding activity of HP1, whereas glutamate substitution mutations, which mimic the charge contributions of phosphorylated serine, have apparently wild-type binding activity. We propose that phosphorylation of HP1 promotes protein-protein interaction between HP1 and target binding proteins in heterochromatin.

Heterochromatin-associated protein 1 (HP1) is a nonhistone chromosomal protein with a dose-dependent effect on heterochromatin mediated position-effect silencing. It is multiply phosphorylated in vivo. Hyperphosphorylation of HP1 is correlated with heterochromatin assembly. We report here that HP1 is phosphorylated by casein kinase II in vivo at three serine residues located at the N and C termini of the protein. Alanine substitution mutations in the casein kinase II target phosphorylation sites dramatically reduce the heterochromatin binding activity of HP1, whereas glutamate substitution mutations, which mimic the charge contributions of phosphorylated serine, have apparently wild-type binding activity. We propose that phosphorylation of HP1 promotes protein-protein interaction between HP1 and target binding proteins in heterochromatin.
In higher eukaryotes, chromosomes are composed of euchromatin and heterochromatin. Heterochromatin is distinguished from euchromatin in that it stays condensed throughout the cell cycle (1). In addition, it replicates late in S phase, is enriched in repetitive DNA, and is relatively poor in classical genes (2)(3)(4).
In Drosophila, when a euchromatic gene is placed next to or within heterochromatin by chromosomal rearrangement or transposition, the euchromatic gene usually undergoes cellspecific silencing, called "position-effect variegation" (5). This process indicates that heterochromatin interferes with euchromatic gene expression, which biochemical data suggest is caused by the acquisition of a distinct chromatin structure with reduced accessibility to DNA-binding proteins (6, 7). More than 50 different position-effect variegation regulators have been identified, including genes encoding chromatin proteins and protein modifiers (8). One such regulator encodes heterochromatin-associated protein 1 (HP1), 1 a nonhistone chromosomal protein enriched in the pericentric heterochromatin of interphase nuclei (9,10). HP1 exerts dosage-dependent effects on position-effect variegation (11,12). HP1 localization during the cell cycle is complex, perhaps reflecting the changes in chromosome structures that accompany the cell cycle (13).
Current models suggest that HP1 functions as part of a chromosomal protein complex (14). Proteins thought to bind to HP1-like proteins include transcriptional intermediary factors (15), lamin B receptors (16), and origin recognition complex proteins (17,18).
HP1 is multiply phosphorylated in vivo (19). Using twodimensional gel electrophoresis, HP1 can be resolved into as many as eight charged isoforms. HP1 phosphorylation occurs predominantly at serines and threonines, and increased phosphorylation of HP1 is correlated with heterochromatin assembly during development. Phosphorylation may be correlated with the oligomeric state of HP1 in vivo (18). Phosphorylation of chromosomal proteins is implicated in the regulation of several nuclear functions. For example, histones are multiply phosphorylated during mitotic and developmentally programmed chromatin condensation (20), specific forms of phosphorylated histone H1 are correlated with specific heterochromatic satellite DNA sequences (21), and several transcription factors are regulated by phosphorylation (22).
As a first step toward defining the role of HP1 phosphorylation in heterochromatin assembly and position-effect variegation silencing, we mapped three of the phosphorylation sites at the N-and C-terminal domains of HP1. We present biochemical evidence for casein kinase II (CKII) phosphorylation of HP1 in vitro and in vivo. Alanine substitution mutation in the Nterminal CKII target site dramatically reduces heterochromatin binding activity of HP1, whereas glutamate substitutions mutations at either or both of the N-and C-terminal sites have apparently wild-type binding activity, suggesting that CKII phosphorylation is required for heterochromatin binding. We propose that phosphorylation of HP1 promotes protein-protein interaction between HP1 and target binding proteins within heterochromatin.

EXPERIMENTAL PROCEDURES
Fly Culture and Tissue Preparation-Fly stocks were maintained at room temperature on standard cornmeal-yeast-sucrose-agar medium containing methylparaben as a mold inhibitor. Third-instar larvae were collected immediately before dissection and kept on ice until dissection.
Expression and Purification of Recombinant HP1-A XbaI-BamHI fragment containing Drosophila HP1 cDNA was cloned into expression vector pET11a, and the recombinant plasmid was transformed into Escherichia coli BL21(DE3) cells. A single colony of transformed cells was grown in LB medium until A 600 ϭ 0.6 -1.0. Expression of HP1 was induced by 1 mM isopropyl-1-thio-␤-D-galactopyranoside at 30°C for 2-3 h. Cells were spun down at 5,000 ϫ g for 5 min in 4°C, and the cell pellet was frozen and stored at Ϫ70°C. For protein purification, the pellet was thawed on ice and suspended in 40 ml of lysis buffer (100 mM Tris-HCl, pH 7.3, 4 mM EDTA, 0.4 mM EGTA, 4 mM dithiothreitol, and 1 mM phenylmethylsulfonyl fluoride). All subsequent steps were done on ice or at 4°C unless indicated otherwise. 24 mg of lysozyme was added to the sample and incubated on ice for 20 min. Cells were lysed either by freezing and thawing or by sonication. NaCl was added to a final concentration of 0.3 M to get maximum HP1 solubilization. After centrifugation at 18,000 rpm for 30 min in a Sorvall SS-34 rotor, the supernatant was diluted 3-fold and applied to a DEAE-Sepharose col-umn. The DEAE column was eluted with 0.4 M NaCl. Eluted fractions containing HP1 were pooled, diluted 4-fold, and applied to a 6-ml Resource Q FPLC (Amersham Pharmacia Biotech FPLC system) column. The Resource Q column was developed with a 0.1-0.4 M NaCl gradient, and HP1 was eluted in the 0.2-0.25 M fractions. These fractions were pooled and loaded on an S-200 Sephacryl FPLC gel filtration column. HP1 was collected in an elution volume corresponding to a molecular mass of ϳ40 kDa. HP1-containing fractions were pooled again and applied to a 1-ml Mono Q FPLC column. The Mono Q column was developed with a 0.1-0.4 M NaCl gradient, and HP1 was purified as a single sharp, symmetrical peak at ϳ0.25 M NaCl. At each chromatographic step, fractions containing HP1 were determined by 12% SDS-PAGE and Coomassie Brilliant Blue staining. Relative purification at each step is shown in Fig. 1.
Preparation of Drosophila Nuclear Extract-Five g of frozen 0 -6-hold embryos (gift of Dr. S. C. R. Elgin) were used to make the nuclear extract. Drosophila embryo nuclei were prepared according to Wu et al. (23). The nuclear extract was made as described (9), with modifications. Briefly, nuclei were suspended in 4 ml of extraction buffer (10 mM Tris-HCl, pH 7.4, 1 mM EDTA, 1 mM ␤-mecaptoethanol, 0.25 M sucrose, 1 mM phenylmethylsulfonyl fluoride, 0.4% Triton X-100) and homogenized at 4°C with a glass-Teflon homogenizer. Homogenate was centrifuged in an Eppendorf 5415 C centrifuge at top speed for 10 min in 4°C. The supernatant was aliquotted and stored at Ϫ70°C as nuclear extract. Protein concentration of the extract was found to be about 1 mg/ml with a BCA protein assay (Pierce).
In Vitro Phosphorylation Assay-In a 20-l reaction mixture, purified recombinant HP1 (1 g/l) was mixed with 1 l Drosophila embryo nuclear extract and 2 Ci of [␥-32 P]ATP at 100 Ci/mol in 20 mM Tris-HCl, pH 7.4, 10 mM MgCl 2 , and 100 mM KCl or 100 mM NaCl (both salts work equally well). The reaction proceeded at room temperature for 20 min and was stopped by adding 5 l of 5ϫ SDS-PAGE sample buffer and boiling for 3 min. Products were separated by a 12% SDS-PAGE, and radiolabeled proteins were visualized by exposing the gel to x-ray film or imaging using a Molecular Dynamics PhosphorImager.
Two-dimensional Gel Analysis and Western Blotting-[ 32 P]Phosphate metabolic labeling of larval tissue and separation of in vivo labeled HP1 by two-dimensional gel electrophoresis was done as described (19). For analysis of in vitro phosphorylated HP1, protein was precipitated with 10% trichloroacetic acid, washed successively with 100% ethanol and chloroform:methanol (1:1), dried, and resuspended in 80 ml of IEF sample buffer (9.5 M urea, 2% Triton X-100, 5% ␤-mercaptoethanol, 1.6% Bio-Lyte 5-8 ampholytes (Bio-Rad), 0.4% Bio-Lyte 3-10 ampholytes (Bio-Rad)) before being applied to an isoelectric focusing gel. After two-dimensional gel electrophoresis, proteins were transferred to nitrocellulose (Millipore) membrane using a Bio-Rad mini-Trans-Blot electrophoretic transfer cell according to manufacturer's instructions. After transfer, the membrane was blocked with 1% bovine serum albumin (fraction V, Sigma) in TBST buffer (10 mM Tris-HCl, pH 8.0, 150 mM NaCl, 0.05% Tween 20) at room temperature for 30 min. The anti-HP1 serum was a gift of Drs. R. F. Clark and S. C. R. Elgin (Washington University, St. Louis, MO) and was used at a 1:10,000 dilution in TBST buffer. The secondary antibody was an anti-rabbit IgG-alkaline phosphatase conjugate (Promega) and was used at a 1:7,500 dilution in TBST buffer. Detection was with 5-bromo-4-chloro-3-indoyl phosphate and nitro blue tetrazolium (Promega) as described (24). To detect radiolabeled protein, Western blots were either subjected to autoradiography using x-ray film (Kodak XAR-5) or imaged using a Molecular Dynamics PhosphorImager.
Trypsin Digestion and Radio-peptide Mapping-In situ trypsin digestion of protein immobilized on nitrocellulose membrane was done according to Fernandez et al. (25). Briefly, the radiolabeled proteins were transferred electrophoretically to nitrocellulose membrane after SDS-PAGE. Radiolabeled HP1 was visualized by Western blotting and autoradiography. The region of membrane containing immunolocalized HP1 was excised with a surgical blade and blocked in 1.0 ml of 0.5% polyvinylpyrrolidone, M r 40,000 (PVP-40, Sigma), 100 mM acetic acid (w/v) in a 1.5-ml Eppendorf tube at 37°C for 30 min. The sample was washed six times with 1 ml of Milli-Q water, cut into approximately 1 x 1-mm squares, and returned to the same tube. A solution of 0.1 M NH 4 HCO 3 and 2 M urea was added to just immerse the membrane strips. Trypsin (0.5 g/g of substrate) was added, and the digestion was incubated overnight at 37°C. Following the digestion, the sample was centrifuged at top speed in an Eppendorf centrifuge for 5 min. The supernatant was stored at Ϫ20°C before peptide mapping. Tryptic peptide mapping was done on a 40% PAGE under alkaline conditions as described (26,27). The gel was dried and subjected to autoradiography or PhosphorImager analysis to visualize the phosphopeptides.
Sequencing of Radiolabeled Tryptic Peptides and Identification of Radioactive Amino Acids-Trypic phosphopeptide bands on the dried PAGE gel were excised with a surgical blade. Peptide was eluted from the gel slices with Milli-Q water. Peptides in the extract were purified and desalted with a C-18 reverse phase HPLC column (130A Separation System, Applied Biosystems) and subjected to protein sequencing (477A Protein Sequencer, Applied Biosystems). Cleaved phenylthiohydantoin derivatives from each cycle were collected, and radioactive amino acid residues were determined by: 1) counting in a scintillation counter (1500 Tri-Carb liquid scintillation analyzer, Packard Instrument Co.) for the radioactivity and 2) concentrating to Ͻ10 l and spotting onto a 0.1-mm cellulose thin layer electrophoresis plate (EM Separations) for autoradiography.
Anti-CKII Immunodepletion of Extracts-Antibody depletion of CKII was done according to Birnbaum et al. (28) with modifications. 5 l of nuclear extract was incubated with 4 l of rabbit antiserum against native Drosophila CKII, 16 l of NET buffer (140 mM NaCl, 50 mM Tris-HCl, pH 8.0, 5 mM EDTA, and 0.05% Nonidet P-40), and 25 g of bovine serum albumin at 4°C for 1 h. 25 l of protein A-acrylic beads suspension (Sigma) was added, and the mixture was incubated at 4°C for an additional 20 min. Following a brief centrifugation, the supernatant was used in the in vitro phosphorylation assay. As a control, 5 l of the extract was treated with rabbit nonimmune serum (no preimmune serum remains), 2 and the sample was also used in the in vitro phosphorylation assay.
Site-directed Mutagenesis and Construction of the Transgene Expression Vector-Site-directed mutagenesis was performed using the Transformer site-directed mutagenesis kit (CLONTECH) according to manufacturer's instructions. Mutagenesis was done in a pBluescript SK(ϩ) plasmid (Stratagene) with Drosophila HP1 cDNA inserted between the XbaI and BamHI sites. Mutant cDNA was then excised with SacI and BglII and cloned into vector SP1 (29). A KpnI fragment from the SP1 vector was cloned into transformation vector pV␤206 in which HP1 cDNA is fused downstream of and in frame with E. coli lacZ under the Drosophila Hsp70 heat-shock promoter (29), inserted in the P-element vector, pYC1.8 (30).
Expression and Localization of ␤-Galactosidase Fusion Protein-Germline transformation was performed by injecting v 36F ;ry 506 embryos with each of the constructs together with the helper plasmid p25.7wc (31), essentially as described (32). G o survivors were mated to v 36F ;ry 506 flies. F 1 adults were screened for rescue of the vermilion eye color. Somatic transformation was performed essentially as described (33,34). Host strain in both cases was a v 36F ;ry 506 stock. Third-instar larvae were collected from each of the germline and somatic transformants carrying the pv␤206 mutant fusion constructs, heat-shocked at 37°C for 30 min, and recovered at room temperature for 1 h. Salivary glands were dissected and stained with X-gal (5-bromo-4-chloro-3-indolyl ␤-Dgalactopyranoside) as described (35). Stained tissue was placed on a microscope slide and mounted in 95% glycerol, 5% phosphate-buffered saline.

Drosophila Embryo Nuclear Extract Phosphorylates Recombinant HP1 in Vitro-HP1 is multiply phosphorylated in vivo,
and hyperphosphorylation is correlated with heterochromatin assembly (19). As a first step toward understanding the functional significance of HP1 phosphorylation, we mapped phosphorylation sites in HP1. Our strategy was to use recombinant HP1 (rHP1) as a substrate for phosphorylation by Drosophila nuclear extract in vitro and then map the sites of in vitro phosphorylation corresponding to sites used in vivo. Drosophila HP1 cDNA was expressed in E. coli and purified to Ͼ95% homogeneity (Fig. 1). To determine whether rHP1 is phosphorylated in bacteria, we compared its mobility in two-dimensional gel electrophoresis to that of HP1 from Drosophila (dHP1). Multiple dHP1 isoforms are detected in Western blots of total Drosophila protein after two-dimensional gel electrophoresis (Ref. 19; Fig. 2A, top). Similar Western blots of rHP1 revealed a single major HP1 isoform ( Fig. 2A, middle). When rHP1 is added to total Drosophila protein and co-electrophoresed, the rHP1 migrates at the extreme basic end of the dHP1 pattern ( Fig. 2A, bottom), consistent with unmodified HP1.
Hyperphosphorylated isoforms of HP1 begin to appear by 2 h of embryonic development (19). Therefore, we used a nuclear extract from 0 -6-h Drosophila embryos to phosphorylate rHP1 in vitro. When rHP1 is added to nuclear extract in the presence of [ 32 P]ATP, rHP1 is efficiently radiolabeled. Co-electrophoresis with total Drosophila protein demonstrates that radiola-beled rHP1 co-migrates with the second and third most basic isoforms of dHP1 (Fig. 2B), consistent with mono-and diphosphorylated protein. To verify that sites of rHP1 phosphorylation by embryo extract correspond to sites of dHP1 phosphorylation in vivo, the major tryptic phosphopeptides of in vitro labeled rHP1 were compared with those of dHP1 metabolically labeled with [ 32 P]orthophosphate. In vitro phosphorylated rHP1 was fractionated by SDS-PAGE and treated with trypsin. Separately, total protein from larval tissue metabolically labeled with [ 32 P]orthophosphate was fractionated by two-dimensional gel electrophoresis, and dHP1 protein was digested with trypsin. Tryptic peptides from both preparations were resolved electrophoretically on a 40% alkaline polyacrylamide gel. Three major radioactive bands were common to both preparations (Fig. 2C). Thus, these major tryptic phosphopeptides contain targets of phosphorylation used in vivo.
HP1 Is Phosphorylated by CKII-The fact that embryo extract can efficiently phosphorylate rHP1 at sites used in vivo allowed us to map the residues phosphorylated by the extract. The major tryptic peptides from in vitro phosphorylated rHP1 (peptides 1, 2, and 3 in Fig. 2C) were size-fractionated by 40% alkaline PAGE, and each phosphopeptide was separately eluted from the gel, desalted and purified by reverse phase HPLC, and subjected to amino acid sequencing. As each amino acid was determined, the eluted amino acid derivative was assayed for radioactivity. Results are summarized in Fig. 2D. Peptides 1 and 2 are fragments of the HP1 N terminus and contain the same phosphorylated serine residue, with peptide 2 having one more undigested lysine at the N terminus. Peptide 3 is a C-terminal fragment containing a major phosphoserine and a minor one three residues away on its N-terminal side. The two most terminal serines occur within CKII recognition motifs, which is S(T)XXE(D). These are the only consensus CKII site motifs in HP1. In peptide 3, the N-terminal serine resembles a CKII target only after its C-terminal partner is phosphorylated and becomes acidic.
To confirm the identity of the HP1 kinase activity in embryo extract, we tested the sensitivity of the kinase to inhibitors and stimulators of CKII. rHP1 phosphorylation by embryo extract is stimulated by spermine and inhibited by heparin (Fig. 3). Both of these effects are also seen with CKII (36,37). As further confirmation, we tested whether antibody to Drosophila CKII (a gift of Dr. C. V. C. Glover III) inhibited in vitro phosphorylation. As shown in Fig. 3, the CKII antisera almost completely eliminates HP1 phosphorylation by embryo extract. Taken together, the results strongly implicate CKII in HP1 phosphorylation in vitro and in vivo.
Mutations in CKII Sites Affect Heterochromatin Binding-To test the functional significance of HP1 phosphorylation, the serines in the CKII target sites were replaced by either alanine (to prevent phosphorylation) or glutamate (to mimic the spatial and charge contributions of phosphorylated serine; Refs. 38 and 39). Five mutations were assayed (Fig. 4): replacement of the N-terminal serine by alanine (S15A) or by glutamate (S15E), replacement of the very C-terminal serine by alanine (S202A) or by glutamate (S202E), and a combination of both Ser 3 Ala mutations (S15A,S202A). The mutant HP1 cDNA were fused downstream from, and in frame with, E. coli lacZ, and the fusion cDNA was placed under the control of the Drosophila Hsp70 heat shock promoter. P-element-mediated germline transformations were generated for the two glutamate substituted mutants. Transgenic larvae were subjected to heat shock, and larval polytene tissue was stained with X-gal to localize the fusion protein. ␤-Galactosidase alone is a cytoplasmic protein when expressed in Drosophila cells (Fig. 4a). As reported previously (29,40), full-length HP1⅐␤-galactosidase fusion protein targets ␤-galactosidase activity to the nucleus and decorates the heterochromatin with ␤-galactosidase; this results in blue-staining nuclei with X-gal, with one or two intensely staining dark blue spots corresponding to the pericentric heterochromatin (Fig. 4c). For both the S15E and S202E mutations, X-gal staining showed that these two mutant HP1 fusion proteins resulted in nuclear localization and heterochromatin binding indistinguishable from wild-type HP1 fusions (Compare Fig. 4, e and g-c; see also Refs. 29 and 40).
For the S15A, S202A, and S15A,S202A mutations, comparable efforts to obtain germline transformants resulted in no transgenic lines (occasionally, transformed phaerate adults were observed, but these failed to hatch or died shortly after hatching). To assay fusion protein targeting for these mutations, we resorted to somatic transformation. Plasmid DNA injected into early embryos stably endures in somatic tissues through the third-instar larval stage, displaying correct tissuespecific expression (33,34). We had employed somatic transformation in previous studies to assay HP1⅐␤-galactosidase fusions (29,40). Somatically transformed third-instar larvae were heat-shocked, and polytene tissues were dissected out and stained with X-gal. By this assay, the S202A mutant appeared similar to the wild type (Fig. 4f), but the S15A mutant showed decreased heterochromatin binding activity visible only in highly expressing cells (Fig. 4d). Only in overstained nuclei did we observe the darkly staining spots that represent heterochromatin binding. For the double mutation (S15A,S202A), the loss of heterochromatin targeting was even more dramatic. Although the fusion protein still appeared to concentrate in the nuclei, it no longer bound to heterochromatin (Fig. 4h). These results demonstrate a requirement for CKII phosphorylation for efficient heterochromatin binding.

Drosophila HP1 Is Phosphorylated by CKII in Vivo-
The methods we used to identify HP1 phosphorylation sites involved direct comparison of the in vivo and in vitro tryptic peptide map by high concentration PAGE, rHP1 phosphopeptide sequencing, and radioactivity detection of each amino acid derivative. This strategy has been used to identify phosphorylation sites on other proteins (26,27,41,42). For all three sites common to phosphorylated rHP1 and dHP1, the targets are good fits to CKII consensus motifs, which, together with the sensitivity of rHP1 phosphorylation to spermine, heparin, and anti-Drosophila CKII serum, strongly suggests that HP1 is a substrate for CKII. CKII is an ubiquitous cyclic nucleotideindependent protein kinase that appears not to directly mediate known signaling pathways (43). CKII activity has been found to increase in response to some mitogens, and its substrates include a number of transcription factors involved in growth control (44). Because CKII is found both in the nucleus and the cytoplasm (36), and because we found that alanine substitution had no effect on nuclear targeting, HP1 phosphorylation by CKII could occur in either compartment.
CKII consensus target sites are found at the N and/or C terminus of HP1 homologs from Drosophila virilis, Schizosaccaromyces pombe, mealybug, mouse, and human. Not all HP1 homologs have CKII targets at both ends (some have neither), but in several such cases the homologous position is occupied by glutamate. Little or nothing is known about the functional homology between Drosophila melanogaster HP1 and its structural homologs in other species, but such apparent structural conservation suggests functional conservation. Nevertheless, the data presented here showing that CKII phosphorylation is required for efficient heterochromatin targeting by the unique D. melanogaster HP1 suggest that such structural conservation is likely to be functionally significant.
The only detailed structural information for any HP1 homo-  (29) with wild-type CKII sites, showing heterochromatin binding; d, S15A⅐␤-galactosidase fusion protein, showing nuclei from highly expressing cells in which reduced heterochromatin binding can sometimes be seen; e, S15E⅐␤-galactosidase fusion protein; f, S202A⅐␤-galactosidase fusion protein; g, S202E⅐␤-galactosidase fusion protein; h, S15A,S202A⅐␤-galactosidase fusion protein. In the nuclei of highly expressing cells, a faint spot of staining is sometimes seen (arrowhead in the right nucleus).
log is a solution NMR peptide structure based on the N-terminal chromo domain of a single mouse HP1 homolog (45), and the sequences corresponding to the targets of CKII lie outside of the solved structure. So far, we have found no effect of CKII phosphorylation on HP1 multimerization in solution. 3 Although two of the CKII sites occur within a previously identified nuclear targeting domain (40), we observed no impairment in nuclear targeting for any of the CKII site mutant fusions. CKII phosphorylation could contribute to heterochromatin binding by HP1 by promoting a conformational shift that permits 1) additional kinases to phosphorylate internal targets in, for example, the HP1 linker region between the chromo domains; or 2) the exposure of sites for protein-protein interactions. Either of these results could facilitate heterochromatin assembly. As previously noted (19), this interval is serine/ threonine-rich and includes two consensus targets for protein kinase A and one for protein kinase C.
Phosphorylation on Both the N-and the C-terminal CKII Sites Is Required for Heterochromatin Binding-Although the Ser 3 Ala mutation on the C-terminal site did not discernibly alter the heterochromatin binding activity of the mutant fusion protein, the Ser 3 Ala mutation on the N-terminal site conspicuously reduces heterochromatin binding. The double Ser 3 Ala mutation (S15A,S202A) almost completely eliminated heterochromatin binding, although the protein could still get into the nucleus. The double mutant appeared to have a generally more severe effect. However, care should be taken in interpreting quantitative differences, because levels of fusion protein expression vary from cell to cell in these assays. Although the effect of the single C-terminal substitution was not detectable by the X-gal staining method, it is possible that each mutation exerts some effect on HP1 heterochromatin binding activity because the combined mutations had the most dramatic effect on heterochromatin binding. Although there are two CKII sites at the C terminus of HP1, we chose to mutate the more downstream site. The upstream site is dependent on the phosphorylation of the downstream serine, so when we mutated the first serine to alanine, we also disabled the second one as a CKII target. Thus, in our assay, any effect attributable to the downstream serine could also reflect a requirement for phosphorylation of the upstream serine. Experiments are in progress to identify additional sites of HP1 phosphorylation and to test their role in HP1 localization and silencing activity.
Although transformants were recovered with Ser 3 Glu mutations in the N or C terminus, we were unable to recover germline transformants with Ser 3 Ala mutations. The significance of this finding is unclear, but it may represent a kind of "dominant negative" phenotype (46). In the absence of heat shock, basal levels of wild-type HP1⅐␤-galactosidase fusion protein are not toxic (29,40), although such transgenic lines are not as healthy as wild-type flies. 3 Mutant HP1 fusion protein, on the other hand, may be toxic at low basal levels. A reasonable speculation is that the nonphosphorylated HP1 participates in only some HP1-dependent activities or sequesters heterochromatin factors in an inactive form.
CKII Phosphorylation and HP1 Function-The most basic HP1 isoforms in vivo are phosphorylated at CKII sites. Thus, CKII phosphorylation does not directly account for the hyperphosphorylation that accompanies the appearance of heterochromatin in the early embryonic development. Indeed, it probably accounts for the maternally loaded HP1 isoforms seen in unfertilized oocytes (19). Nevertheless, the mutational analysis shows that CKII phosphorylation is essential for heterochromatin binding.
CKII is an ubiquitous eukaryotic serine/threonine protein kinase that phosphorylates more than 100 substrates, many of which control cell division or signal transduction. These substrates include a striking number of nuclear proteins involved in DNA replication and transcription (47). CKII modifies protein-DNA binding (48 -51) and protein-protein interaction (52,53). In Drosophila, CKII is present in both the cytoplasmic and nuclear compartments. CKII phosphorylation enhances the DNA binding activity of the engrailed protein (51) and modulates Antennapedia activity (39) and dorsoventral patterning (54). Drosophila DNA topoisomerase II is stimulated by CKII phosphorylation (55). We showed previously that significant HP1 phosphorylation still occurs in vivo in tissues treated with sufficient cycloheximide to block all detectable nascent protein synthesis (19). This turnover of phosphate uncoupled from a new synthesis suggested that HP1 phosphorylation could regulate its chromatin association, an example being the dynamic dissociation and reassociation of HP1 that reportedly takes place during mitosis (13). Alternatively, phosphorylation-dephosphorylation may be regulated during decondensation of heterochromatin to permit DNA replication in late S phase.


extfieldsofstudy: Medicine Biology

text:
Alternatively Spliced EDA Segment Regulates Fibronectin-dependent Cell Cycle Progression and Mitogenic Signal Transduction*

Fibronectin (FN) is comprised of multiple isoforms arising from alternative splicing of a single gene transcript. One of the alternatively spliced segments, EDA, is expressed prominently in embryonic development, malignant transformation, and wound healing. We showed previously that EDA+ FN was more potent than EDA− FN in promoting cell spreading and cell migration because of its enhanced binding affinity to integrin α5β1 (Manabe, R., Oh-e, N., Maeda, T., Fukuda, T., and Sekiguchi, K. (1997) J. Cell Biol. 139, 295–307). In this study, we compared the cell cycle progression and its associated signal transduction events induced by FN isoforms with or without the EDA segment to examine whether the EDA segment modulates the cell proliferative potential of FN. We found that EDA+ FN was more potent than EDA− FN in inducing G1-S phase transition. Inclusion of the EDA segment potentiated the ability of FN to induce expression of cyclin D1, hyperphosphorylation of pRb, and activation of mitogen-activated protein kinase extracellular signal regulated kinase 2 (ERK2). EDA+ FN was also more potent than EDA− FN in promoting FN-mediated tyrosine phosphorylation of p130Cas, but not focal adhesion kinase, which occurred in parallel with the activation of ERK2, suggesting that p130Cas may be involved in activation of ERK2. These results indicated that alternative splicing at the EDA region is a novel mechanism that promotes FN-induced cell cycle progression through up-regulation of integrin-mediated mitogenic signal transduction.

Fibronectins (FNs) 1 are extracellular matrix glycoproteins that play a role in cell adhesion and migration during embryonic development, wound healing, and tumor progression (1). Proteolytic digestion of FN has revealed that FN has several cell binding domains: the central cell binding domain (CCBD), the COOH-terminal heparin binding domain (Hep2), and the CS1 region within the type III connecting segment (IIICS) (2). CCBD is the major cell-adhesive domain of FN and contains the Arg-Gly-Asp (RGD) motif that is recognized by members of the integrin family of cell adhesion receptors including ␣5␤1, ␣v␤1, ␣v␤3, ␣v␤5, ␣v␤6, ␣IIb␤3, and ␣8␤1 (3)(4)(5)(6). There is evidence that FN acts not only as a cell-adhesive substrate but also transduces biochemical signals across the plasma membrane via integrin receptors, thereby regulating cell proliferation, differentiation, and apoptosis (7)(8)(9). Interaction of FN with integrins stimulates tyrosine phosphorylation of several cellular proteins including FAK (10 -12) and p130 Cas (13)(14)(15) and also stimulates activation of Src family protein tyrosine kinases (7) and the extracellular signal-related kinases ERK1 and ERK2 (16 -19). Furthermore, FN induces expression of cyclin D1 and subsequent inactivation of pRb by hyperphosphorylation and thus mediates cell cycle progression through the G 1 phase (20).
FN contains three alternatively spliced segments: EDA, EDB, and IIICS (21)(22)(23)(24). The EDA and EDB segments are each encoded by a single exon, and both comprise an intact type III repeat (22). FNs expressed in fetal and tumor tissues contain a greater percentage of EDA and EDB segments than those expressed in normal adult tissues (25)(26)(27)(28)(29). Increased expression of FNs containing the EDA and/or EDB segments has also been observed during wound healing (30). Because FNs containing the EDA and/or EDB segments are highly expressed in tissues that are populated with cells showing high proliferative and migratory potential, it seems likely that FNs containing these extra segments play an important role in promoting cell proliferation and migration. Despite accumulated evidence for regulated expression of EDA-and/or EDB-containing FNs in vivo, the biological functions of these isoforms are poorly understood (31).
Recently, we found that FN isoforms containing the EDA segment (EDA ϩ FN) were more potent than those lacking the EDA segment (EDA Ϫ FN) in promoting cell adhesion and migration, irrespective of the presence or absence of the EDB segment. The increased cell adhesion and migration were caused by increased binding of EDA ϩ FN to integrin ␣5␤1 (32). In the present study, we investigated whether alternatively spliced EDA and EDB segments regulate the potential of FNs to promote cell proliferation. We found that insertion of the EDA, but not EDB, segment potentiated the ability of FN to promote cell cycle progression. The enhanced cell cycle progression was associated with increased tyrosine phosphorylation of p130 Cas , activation of ERK2, increased expression of cyclin D1, and hyperphosphorylation of pRb. These results indicated that alternative splicing in the EDA region regulates FN-mediated extracellular signals and subsequent cell cycle progression via modulation of binding affinity to integrin ␣5␤1.
viously (32). To construct the expression vector for EDA Ϫ /EDB ϩ FN, the whole inserts of pHCF93B ϩ encoding Val 527 -Arg 1449 including the EDB segment (32) were excised by double digestion with SalI/BamHI and ligated to SalI/BamHI-cleaved pAIFNC, which encodes a full-length FN isoform lacking both EDA and EDB segments. The resulting cDNA expression vector for EDA Ϫ /EDB ϩ FN was designated pAIFNBC. Recombinant FNs were expressed in CHO DG44 cells and purified from the culture supernatants of the stable transfectants by gelatin-affinity chromatography as described previously (32).
Cell Culture-CHO-K1 and CHO DG44 cells were obtained from the Japanese Cancer Research Resources Bank (Tokyo, Japan) and Dr. Lawrence Chasin (Columbia University), respectively. CHO-K1 cells were grown in Ham's F-12 medium supplemented with 10% FBS. CHO DG44 cells deficient in dihydrofolate reductase activity were maintained in ␣-minimal essential medium containing ribonucleosides/deoxyribonucleosides and 10% FBS. Stable transfectants of CHO DG44 cells overexpressing recombinant FNs were maintained in ␣-minimal essential medium without ribonucleosides/deoxyribonucleosides in the presence of 10% FBS (for routine passage) or 1% FN-depleted FBS (for large scale culture to purify recombinant proteins). All cells were maintained under a humidified 5% CO 2 atmosphere at 37°C.
Cell Spreading Assay-Cell spreading assays were performed using 96-well microtiter plates coated with 5 g/ml recombinant FNs and blocked with 1% bovine serum albumin. Amounts of recombinant FNs immobilized on plates were determined by enzyme-linked immunosorbent assay using anti-human FN antiserum or anti-FN mAbs as described previously (32). CHO-K1 cells grown in Ham's F-12 medium containing 10% FBS were trypsin treated and washed with serum-free medium. The cells were plated at a density of 3 ϫ 10 4 cells/well in serum-free medium and incubated at 37°C for 40 min. Nonadherent cells were removed by washing with serum-free medium, and attached cells were fixed with 3.7% formaldehyde and then stained with Giemsa. Cells adopting a well spread morphology (i.e. cells that had become flattened with the long axis more than twice the diameter of the nucleus) were counted/mm 2 .
Induction of G 1 -S Phase Transition in Synchronized Cells-CHO-K1 cells were grown to confluence and synchronized at G 0 by serum starvation for 48 h in Ham's F-12 supplemented with 0.2% FN-depleted FBS, 50 g/ml streptomycin, and 50 units/ml penicillin (starvation medium) according to Orren et al. (33). The cells were dissociated with phosphate-buffered saline containing 0.2% trypsin and 2 mM EDTA, and trypsin treatment was stopped by adding soybean trypsin inhibitor (1 mg/ml). The cells were washed with serum-free medium, incubated in suspension in starvation medium for 90 min at 37°C, and seeded on plates that had been precoated with 5 g/ml recombinant FNs or 20 g/ml poly-L-lysine and then blocked with 1% bovine serum albumin. The amounts of immobilized recombinant FNs on the plates were determined by enzyme-linked immunosorbent assay using anti-human FN antiserum to confirm the equality of adsorbed FNs.
Determination of S Phase Entry-Quiescent cells were kept in suspension for 90 min and then plated at a density of 2 ϫ 10 4 cells/well in starvation medium. [ 3 H]Thymidine (5 Ci/ml) was added 12-24 h after cell plating. The cells were washed three times with serum-free medium at the indicated time points, fixed with ice-cold 10% trichloroacetic acid for 1 h, and washed once with 10% trichloroacetic acid. The acidinsoluble precipitates were dissolved with 0.1% SDS and 0.2 N NaOH, neutralized with 0.2 N HCl, and transferred to vials containing scintillation fluid. Radioactivities in the precipitates were determined with an Aloka LSC-3500 scintillation counter. To determine the percentage of cells entering S phase, bromodeoxyuridine was added to the cultures 12-24 h after plating, and the cells were incubated with anti-bromode-oxyuridine mAb followed by visualization of bound mAb with 3,3Јdiaminobenzidine tetrahydrochloride according to the manufacturer's instructions (Amersham Pharmacia Biotech).
For immunoprecipitation of FAK, cell lysates were incubated with protein A-conjugated anti-FAK mAb 2A7 for 5 h. For immunoprecipitation of p130 Cas , cell lysates were incubated with anti-p130 Cas antibody for 1 h followed by incubation with protein G-Sepharose (Amersham Pharmacia Biotech) for 4 h. The immune complexes were precipitated by brief centrifugation and washed four times with lysis buffer. The immunoprecipitates were separated on 8% polyacrylamide gels to determine the levels of tyrosine phosphorylation and the amounts of precipitated proteins as described above.

EDA but Not EDB Regulates Cell Adhesive Functions of FN-
We showed previously that the presence or absence of the EDB segment did not affect the ability of EDA ϩ FN isoforms to promote cell adhesion and spreading (32), although we could not exclude the possibility that inclusion of the EDB segment alone might potentiate the cell-adhesive activity of FN. To explore this possibility, we produced a recombinant FN isoform containing the EDB but not the EDA segment (designated rFN(BC); see Fig. 1) and compared its cell attachment and spreading activities with those of rFN(C), rFN(AC) and rFN-(BAC). CHO-K1 cells were used in these assays because they have long been used as a standard model in cell cycle analysis and are more easily synchronized at the G 0 state than HT1080 cells, which were used in the previous study (32). Significant differences were observed in both cell attachment and spreading among the four different FN isoforms, EDA ϩ FNs (i.e. rFN(AC) and rFN(BAC)) being more potent in promoting cell attachment than EDA Ϫ FNs (i.e. rFN(C) and rFN(BC)) ( Fig.  2A). EDA ϩ FNs were also more than twice as effective as EDA Ϫ FNs in inducing cell spreading (Fig. 2B). No significant differences were observed in cell spreading activity between rFN(AC) and rFN(BAC) or between rFN(C) and rFN(BC) (Fig.  2B), indicating that the EDB segment is not involved in regulation of the cell-adhesive activity of FN. Because CHO-K1 cells adhere to FN-coated substrates predominantly via integrin ␣5␤1 (34) and the increased cell-adhesive activity of EDA ϩ FNs is the result of enhanced binding of CCBD to integrin ␣5␤1 (32), the integrin binding activity of EDB ϩ FN was compared with that of EDB Ϫ FN. There were no significant differences in integrin ␣5␤1 binding between rFN(C) and rFN(BC), although rFN(AC) was more than twice as active as rFN(C) and rFN(BC) in integrin ␣5␤1 binding (Fig. 3). These results support the previous conclusion that the cell-adhesive functions of FN are specifically regulated by insertion of the EDA but not the EDB segment.
EDA ϩ FN Is More Potent than EDA Ϫ FN in Promoting Cell Cycle Progression-The in vivo expression patterns of EDA ϩ FN suggested a potential role of EDA ϩ FN in cell proliferation (31). To investigate whether insertion of the EDA and/or EDB segment affects the ability of FN to promote cell proliferation, CHO-K1 cells were synchronized at the G 0 phase by serum starvation, incubated in suspension for 90 min, and replated on substrates precoated with different FN isoforms to stimulate the cells to reenter the cell cycle. To confirm that the effects of FN isoforms on cell proliferation were primarily mediated via integrins, the cells were also plated on the substrate coated with poly-L-lysine, which mediates integrin-independent cell adhesion. All recombinant FNs significantly promoted entry into S phase when compared with poly-L-lysine (Fig. 4). The levels of DNA synthesis were different among the FN isoforms; rFN(AC) and rFN(BAC) were more potent than rFN(C) and rFN(BC) in inducing DNA synthesis. No significant differences were observed between rFN(AC) and rFN(BAC) or between rFN(C) and rFN(BC). Labeling of cells with bromodeoxyuridine showed that more than 70% of the plated cells entered the S phase by 24 h after replating onto the FN-coated substrata (data not shown). These results indicated that EDA ϩ FN has a greater potential than EDA Ϫ FN to induce cell proliferation irrespective of the presence or absence of the EDB segment.
EDA Segment Up-regulates Cell Cycle-associated Signal Transduction-It has been shown that pRb plays a key role in the G 1 -S phase transition. Inactivation of pRb by hyperphosphorylation occurs at the restriction point in G 1 as the cell enters the S phase (35). We investigated whether the presence or absence of the EDA segment could modulate FN-induced phosphorylation of pRb. Quiescent cells were trypsin treated, held in suspension to reset anchorage-dependent signals, and replated onto dishes coated with rFN(C), rFN(AC) or poly-Llysine. Hypo-and hyperphosphorylated forms of pRb were distinguished by the difference in their mobility on SDS-polyacrylamide gel electrophoresis. Hyperphosphorylation of pRb was induced on the substrates coated with rFN(C) and rFN(AC) compared with those with poly-L-lysine (Fig. 5, upper panel). The level of pRb hyperphosphorylation was higher with cells plated on rFN(AC) than on rFN(C) through 16 -24 h after plating, indicating that the increased cell cycle progression driven by EDA ϩ FN was associated with the increased hyperphosphorylation of pRb. Although pRb hyperphosphorylation was detectable even on poly-L-lysine upon prolonged incubation, this could be the result of deposition of endogenous FN by CHO cells. Integrin ␣5␤1 was purified from human placenta and reconstituted in phosphatidylcholine liposomes as described under "Experimental Procedures." The integrin ␣5␤1-liposomes were added to microtiter plates precoated with 20 g/ml of different FN isoforms and incubated for 6 h at room temperature. Quantities of bound integrin ␣5␤1-liposomes are expressed as percentages of the total input radioactivity after subtraction of the radioactivity bound to plates coated only with bovine serum albumin. Each bar represents the mean S.D. (n ϭ 6). pRb has been shown to be phosphorylated by cyclin D1⅐cdk4/6 complexes, with concomitant accumulation of cyclin D1 during the G 1 phase (35). Therefore, we examined whether the level of expression of cyclin D1 was also modulated by the EDA segment upon cell adhesion onto FN-coated substrates. When cells were plated onto substrates coated with recombinant FNs with or without the EDA segment, the expression of cyclin D1 was significantly induced by 24 h after replating, although no obvious induction of cyclin D1 expression was observed on the poly-L-lysine-coated substrate (Fig. 5, lower panel). The expression of cyclin D1 was more pronounced with cells plated onto rFN(AC) than onto rFN(C) with a time course profile similar to that of pRb hyperphosphorylation. These results indicated that expression of cyclin D1 and phosphorylation of pRb were coordinately regulated by the EDA segment upon cell adhesion to FN-coated substrates.
EDA Segment Enhances FN-dependent MAP Kinase Activation-Because activation of MAP kinase has been shown to induce expression of cyclin D1 and subsequent hyperphospho-rylation of pRb (36), we investigated whether ERK2 MAP kinase was stimulated differently by EDA ϩ and EDA Ϫ FNs. When quiescent cells were detached and either held in suspension or replated on dishes coated with rFN(C), rFN(AC), or poly-L-lysine, the activated form of ERK2 was detectable as early as 10 min after replating and persisted for over 60 min (Fig. 6). In contrast, no significant activation was observed in cells kept in suspension or replated on the substratum coated with poly-L-lysine. Activation of ERK2 was more prominent on surfaces coated with rFN(AC) than on those coated with rFN(C), indicating that the EDA segment potentiates the ability of FN to activate MAP kinases via integrin-dependent signaling and thereby promotes cell cycle progression.
EDA Increases FN-induced Tyrosine Phosphorylation of p130 Cas -Protein tyrosine phosphorylation triggered by integrin-mediated cell-matrix interaction has been shown to be caused by activation of protein tyrosine kinases including the Src family kinases, FAK and FAK-related kinases (7). These kinases phosphorylate tyrosine residues of their specific substrate proteins, providing binding sites for SH2-containing molecules that are involved in cell proliferation (7). We examined whether the EDA segment modulates the ability of FNs to stimulate tyrosine phosphorylation of cellular proteins. In parallel with the analysis of FN-dependent activation of ERK2, quiescent cells were kept in suspension or replated on dishes precoated with rFN(C), rFN(AC), or poly-L-lysine and incubated for different periods. As shown in Fig. 7A, proteins migrating at positions corresponding to 150, 130, and 80 -60 kDa were tyrosine phosphorylated irrespective of the type of substrate. Among these, tyrosine phosphorylation of the 130-kDa protein was significantly induced in cells plated on substrates coated with recombinant FNs but not in cells either plated on poly-L-lysine-coated substrates or kept in suspension. The kinetics of tyrosine phosphorylation of the 130-kDa protein were similar to those of ERK2 activation (Fig. 6). The 130-kDa protein was tyrosine-phosphorylated at 10 min after replating on FN-coated surfaces, and the levels of tyrosine phosphorylation persisted for more than 60 min. The 130-kDa protein was more prominently tyrosine-phosphorylated in cells on rFN(AC) than those plated on rFN(C).
Cell adhesion to FN-coated substrates has been shown to induce tyrosine phosphorylation of p125 FAK (10 -12) and p130 Cas (13)(14)(15). Immunoprecipitation with anti-FAK and anti-p130 Cas antibodies showed that p130 Cas was phosphorylated in CHO-K1 cells plated onto FN-coated surfaces, although no significant tyrosine phosphorylation was observed with FAK in the same cells (Fig. 7B, upper panel). These results indicated that the tyrosine-phosphorylated 130-kDa protein was p130 Cas but not FAK. Tyrosine phosphorylation of p130 Cas was more pronounced on the substrates coated with rFN(AC) than on those coated with rFN(C). The enhanced phosphorylation of the Quiescent cells were either held in suspension or plated on substrates coated with 5 g/ml rFN(C) or rFN(AC) or 20 g/ml poly-L-lysine (PLL) in starvation medium for various times. Cells were lysed, and MAP kinase activity was analyzed by a band shift assay. The phosphorylated, activated forms of ERK2 (pp42) display more reduced electrophoretic mobility than the unphosphorylated, nonactivated forms of ERK2 (p42). The reproducibility of the results was confirmed in three separate experiments.
130-kDa protein was not the result of an increase in p130 Cas expression because the amount of immunoprecipitated p130 Cas was decreased rather slightly on rFN(AC) than that on rFN(C) (Fig. 7B, bottom panel). Although we could not exclude the possibility that FAK was tyrosine-phosphorylated at levels below the limits of detection, it is likely that p130 Cas is the main target of protein tyrosine kinases activated by integrin-mediated adhesion onto FN-coated substrates and therefore is involved in activation of ERK2 and subsequent cell cycle progression.

Regulation of FN-mediated Signal
Transduction by the EDA Segment-It has been shown that the EDA segment is included in FN species expressed in embryonic tissues, but is spliced out of the molecule as embryonic development proceeds (28,31). In adults, EDA ϩ FN reappears at sites of tissue injury and inflammation (30,38). Despite the close association of the expression of EDA ϩ FN with tissues populated with cells showing high proliferative potential, the physiological significance of this phenomenon is not known.
In this study, we found that EDA ϩ FN was more potent than EDA Ϫ FN in inducing G 1 -S phase transition. The increased ability of EDA ϩ FN to induce G 1 -S phase transition was closely associated with enhanced ERK2 activation, cyclin D1 expression, and pRb phosphorylation. Given that EDA ϩ FN is more potent than EDA Ϫ FN in promoting cell adhesion through enhanced binding to integrin ␣5␤1 (32) and that ligand ligation of integrin ␣5␤1 promotes DNA synthesis through activation of intercellular signal transduction pathways involving ERK2, cyclin D1, and pRb (1,20), it is conceivable that inclusion of the EDA segment up-regulates FN-dependent mitogenic signal transduction through increased binding of FN to integrin ␣5␤1. Because EDA ϩ FN is expressed preferentially in embryonic and tumor tissues, alternative splicing at the EDA region of FN may be instrumental in supporting the high proliferative potential of cells in these tissues.
FN contains two major integrin binding regions: CCBD and CS-1. CCBD is recognized by integrin ␣5␤1 and ␣v-containing integrins, whereas CS-1 is recognized by integrin ␣4␤1. Besides the distinct binding specificities, the integrin binding activities of these two regions are regulated differently at the level of RNA splicing. Thus, the CS-1 region itself is excluded from the FN molecule by alternative splicing (39 -41), regulating its cell adhesive activity in an all-or-none manner. In contrast, CCBD is encoded by constitutive exons, and its cell adhesive activity is regulated by the alternatively spliced EDA segment in a range of 2ϳ3-fold increments. The difference in the mode of regulation of their activities by alternative splicing appears to reflect the physiological roles of these cell adhesive regions. Integrin ␣5␤1, the major receptor for CCBD, is expressed on a wide variety of cell types and plays a central role in adhesion-dependent signaling events involved in proliferation, migration, and survival of cells (1,9). The interaction of integrin ␣5␤1 with CCBD may provide housekeeping signals that need to be maintained strictly at the minimal level required for cell survival. In contrast, integrin ␣4␤1, the specific receptor for CS-1, is expressed on restricted cell types such as lymphocytes (39,42) and elicits a cell type-specific proliferation signal (43). All-or-none regulation of CS-1 activity by alternative splicing may be suitable for such an auxiliary signal, leaving the basal survival signal from CCBD uncompromised.
The interaction of cells with FN is regulated not only by the alternative splicing at the EDA region but also by the affinity states of integrin ␣5␤1. Integrins exist at different affinity states ranging from inactive to fully activated forms (44). The affinity states of integrins are modulated by cellular stimulation by chemoattractants (44) and interaction with membraneassociated proteins such as IAP (integrin-associated protein) (45). Modulation of the ligand binding affinity of integrin ␣5␤1 has been shown to affect FN-dependent cell proliferation (46) and differentiation (47) by modulation of integrin ␣5␤1-mediated signal transduction. It is conceivable, therefore, that alternative splicing of FN and affinity modulation of integrin ␣5␤1 coordinately regulate the integrin-mediated, extracellular signals required for proliferation, differentiation, and survival of cells.
Role of p130 Cas in FN-mediated Signal Transduction-We found that cell adhesion to FN-coated substrata activates ERK2 without significant tyrosine phosphorylation of FAK. Although there is some evidence suggesting that FAK can activate the ERK signaling pathway in response to cell adhesion (16,17,19), it is unclear whether FAK plays a physiological role in the ERK signaling pathway. In fact, recent studies have shown that the activation of ERK in response to integrin ligation is mediated by the adaptor protein Shc independently of FAK (48). In support of this observation, introduction of a dominant-negative version of FAK did not impair the ERK activation elicited by cell adhesion to FN (49). Our results also showed that p130 Cas , but not FAK, was prominently tyrosinephosphorylated in response to cell adhesion onto FN-coated substrata with kinetics similar to those of ERK2 activation. Although the precise molecular mechanisms of p130 Casdependent activation of ERK remain to be defined, tyrosine phosphorylation of p130 Cas was suggested to be involved in ERK activation by the following observations. First, overexpression of p130 Cas or FN-stimulated tyrosine phosphorylation of p130 Cas promotes the SH2-mediated binding of the adaptor proteins Crk and/or Nck to p130 Cas and subsequent association of Crk and/or Nck with the GDP-GTP exchange protein Sos, which could result in Ras activation (50,51). Second, overexpression of protein tyrosine phosphatase 1B, which can also associate with p130 Cas , resulted in dephosphorylation p130 Cas and inhibition of ERK activation, whereas a protein tyrosine phosphatase 1B mutant unable to bind p130 Cas failed to dephosphorylate p130 Cas and therefore failed to inhibit ERK activation (52). Although these observations seem to support the involvement of p130 Cas in ERK activation, further studies are required to define the role of tyrosine phosphorylation of p130 Cas in EDA-dependent enhancement of cell cycle progression.
Adhesive Function of FN Is Independent of the EDB Segment-Our results showed that the abilities of rFN(BC) to promote cell adhesion, integrin ␣5␤1 binding, and cell cycle progression were the same as those of rFN(C) and were significantly lower than those of rFN(AC), excluding the possibility that inclusion of the EDB segment alone might potentiate the adhesive activity of FN. Adhesive functions of FN are therefore regulated exclusively by alternative splicing at EDA but not EDB. Based on the three-dimensional structure of a recombinant FN fragment consisting of III 7 -III 10 modules (53), we proposed that insertion of the EDA segment alters the global conformation of the FN molecule through rotating the NH 2terminal portion of the FN polypeptide relative to the COOH terminus, thereby resulting in the enhanced binding of EDA ϩ FN to integrin ␣5␤1 because of an increased exposure of the RGD motif in III 10 module and/or local unfolding of the module (32). Failure of the EDB segment to substitute for EDA in altering the global conformation of the FN so as to increase the binding affinity to integrin ␣5␤1 indicates that the conformational activation of FN by insertion of an extra type III module is dependent on the position and/or sequence of the inserted module, making the EDA segment most suited for modulation of the binding affinity to integrin ␣5␤1.
Although the presence or absence of the EDB segment did not affect the cell adhesive activity of FNs, the EDB segment may modulate other biological functions of FN. Expression of EDB ϩ FN in vivo is more restricted than that of EDA ϩ FN (28), suggesting that the EDB segment may have an important function only in highly specific situations such as angiogenesis (54) and lung development (55). Regulated in vivo expression of recombinant FN isoforms differing with respect to the presence or absence of the EDA and EDB segments should provide insight into the physiological and pathological roles of the alternatively spliced segments during development, tumorigenesis, and inflammatory processes.