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• Abdominal Manual Muscle Testing

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Preventive engine cleaning enables you to restore engine parts rather than replacing them, thereby saving vehicle owners on costly parts, such as a new turbocharger ($ 1.350 - 3.100), catalytic converter ($ 600 - 2.000), DPF ($ 600 - 2.000) or EGR valve ($ 370 - 500). These problems result mainly from poor combustion, which stifles the engine. So before replacing your engine parts, try Carbon Cleaning. A dirty engine as the result of carbon deposits is the new threat to vehicle performance. Change Your Oil If necessary, take off the undercover. The good news for us DIYers is that most cars don't have them.

Now it's time to locate the oil filter and drain plug. The vast majority of cars have a bottom-mount screw-on filter. In this case, the plug and filter are far apart, meaning I must reposition the drain pan after I drain the oil and before I remove the filter.

With the preliminaries out of the way, it's time to drain the oil out of the engine. It's important to place the drain pan under the drain plug — but not directly under it. The angle of the drain plug will cause the oil to stream out at an angle, so I'm offsetting the pan to that side by several inches. Remove the oil filler cap (the oil will flow out more smoothly and quickly with the cap removed). It's time to put the gloves on. You can usually remove the drain plug with a common end wrench and a bit of muscle.

Go slowly as you remove the drain plug and keep your hands away from the expected path of the oil. It's going to come out quick and warm. This is another reason why it's best not to change oil when the engine and its oil are piping hot. Inspect and clean the oil drain plug while the rest of the oil is draining. If your drain plug does need a replacement washer, make sure the old one isn't stuck to the engine's oil pan. Tighten the drain plug. Theoretically, there is a torque specification for drain plugs, but they're almost never published in the owner's manual.

Remove the oil filter New filters that are properly installed don't go on terribly tight. But they can be hard to get off later because their sealing gaskets swell over time. Use rags to clean as much oil away as you can, paying special attention to the filter sealing surface.

Install the new filter The last messy step involves smearing a dab of new oil on the new filter's O-ring. Spinning the filter on gently until the O-ring makes first contact with the sealing surface. Generally, oil filters are tightened no more than three-quarters of a turn to a full turn beyond the point where the O-ring first contacts the sealing surface. Consult your manual or the oil filter box to confirm the proper amount.

Abstract Zebrafish SmyD1 is a SET and MYND domain-containing protein that plays an important role in myofiber maturation and muscle contraction. SmyD1 is required for myofibril organization and sarcomere assembly during myofiber maturation. Whole-mount in situ hybridization revealed that smyd1 mRNAs are specifically expressed in skeletal and cardiac muscles in zebrafish embryos. However, it is unknown if smyd1 is expressed in other striated muscles, such as cranial and fin muscles, and moreover, the regulatory elements required for its muscle-specific expression. We report here the analyses of smyd1 expression using smyd1-gfp transgenic zebrafish. Smyd1-gfp transgenic zebrafish were generated using the 5.3-kb smyd1 promoter and its 5′-flanking sequence.

GFP expression was found in the skeletal and cardiac muscles of smyd1-gfp transgenic embryos. GFP expression appeared stronger in slow muscles than fast muscles in transgenic zebrafish larvae.

In addition, GFP expression was also detected in cranial and fin muscles of smyd1-gfp transgenic zebrafish larvae. In situ hybridization confirmed smyd1 mRNA expression in these tissues, suggesting that the expression of the smyd1-gfp transgene recapitulated that of the endogenous smyd1 gene. Deletion analysis revealed that the 0.5-kb sequence in the proximal promoter of smyd1 was essential for its muscle specificity.

Together, these data indicate that smyd1 is specifically expressed in most, if not all, striated muscles, and the muscle specificity is controlled by the 5.3-kb promoter and flanking sequences. Developmental Dynamics 235:3306–3315, 2006. © 2006 Wiley-Liss, Inc. INTRODUCTION The acquisition of muscle cell identity requires invariably induction-dependent changes in gene expression. Increasing evidence indicates that chromatin remodeling through histone modifications plays critical roles in gene expression and muscle cell differentiation (McKinsey et al.,; Rupp et al.,; Zhang et al.,; Lee et al.,; Cirillo and Zaret,; Biron et al., ). Histone acetylation often results in transcriptional activation. In contrast, histone deacetylation leads to repression of gene transcription (McKinsey et al., ).

Histone methylation, on the other hand, is associated with both transcription activation and repression (Santos-Rosa et al., ). Histone methylation is involved in various cellular processes, including cell differentiation, proliferation, and cell transformation (Cerny and Quesenberry,; Sims and Reinberg, ).

Histone methylation is carried out by a novel class of SET domain-containing proteins (Rea et al., ). The first SET domain histone methyltransferase was identified as the mammalian Suv39h1, which methylates histone H3 at Lys 9 (Rea et al., ).

In the past few years, over 50 SET domain-containing proteins have been identified (Kubicek and Jenuwein,; Cheng et al., ). SmyD represents a new family of proteins that contain the two highly conserved SET and MYN D domains (Hamamoto et al.,; Tan et al., ). Smyd3 encodes a histone methyltransferase involved in the proliferation of cancer cells (Hamamoto et al., ).

Smyd1, also known as skm-bop, was originally identified as an unknown gene in the opposite transcriptional orientation from the CD8b gene (Hwang and Gottlieb, ). Smyd1 gene encodes several cDNA isoforms generated by alternative splicing (Hwang and Gottlieb, ). Two of the isoforms ( smyd1a and smyd1b) are specifically expressed in skeletal and cardiac muscles (Hwang and Gottlieb,; Gottlieb et al.,; Phan et al.,; Tan et al., ). Functional analyses in mice and zebrafish demonstrate that SmyD1a and SmyD1b play a key role in development of skeletal and cardiac muscles.

Smyd1 null mutant mice fail to develop a right ventricular chamber and die around embryonic day 10.5 (Gottlieb et al., ). Knockdown of smyd1a and smyd1b expression by antisense oligos results in skeletal and cardiac muscle defects in zebrafish embryos. The SmyD1a and SmyD1b knockdown zebrafish embryos cannot swim and have no heartbeat (Tan et al., ). Close examination reveals that knockdown of smyd1 gene expression disrupts myofibril organization and sarcomere formation in zebrafish embryos (Tan et al., ). The biological activity of SmyD1a or SmyD1b is likely associated with histone modification.

Gottlieb and colleagues have demonstrated that SmyD1a or SmyD1b represses gene transcription in vitro, dependent upon histone deacetylases (HDACs). This finding is consistent with the findings that SmyD1a and SmyD1b proteins contain the two important protein domains MYND and SET domains involved in recruitment of HDAC and histone methylation, respectively (Wang et al.,; Lutterbach et al.,; Gelmetti et al.,; Tan et al., ). SmyD1 is clearly expressed in skeletal and cardiac muscles and plays an important role in skeletal and cardiac muscle differentiation. However, little is known about the expression pattern of smyd1 in other striated muscles, such as cranial and fin muscles, and moreover, the regulatory sequences for the muscle-specific expression. A better understanding of smyd1 expression will provide further insights into SmyD1 functions in muscle cell differentiation. In this study, we have characterized smyd1 expression using transient and transgenic analyses in zebrafish embryos. We have demonstrated that the 5.3-kb smyd1 promoter and 5′-flanking sequence was critical for its muscle-specific expression in skeletal, cardiac, cranial, and fin muscles, and Hh signal appeared to be important for maintaining smyd1 expression in adaxial cells that give rise to embryonic slow muscles.

Together, these data indicate that smyd1 is expressed in most, if not all, striated muscles and that the muscle specificity is controlled by the 5.3-kb promoter and flanking sequences. RESULTS Muscle Specificity of the Zebrafish smyd1 Promoter We have previously shown that smyd1 gene encodes two muscle-specific isoforms, smyd1a and smyd1b, that are generated by alternative splicing. Smyd1a and smyd1b are specifically expressed in skeletal and cardiac muscles in zebrafish embryos (Tan et al., ). To determine the regulatory region for the muscle-specific expression, we first characterized the smyd1 gene promoter and 5′ flanking sequence.

The transcription start site was determined by 5′-rapid amplification of cDNA ends (RACE) at 18 bp upstream of the ATG start codon in exon 1 (Fig. The 5.3-kb DNA sequence including the smyd1 5′-untranslated region (UTR), promoter, and its flanking sequence was cloned upstream of a green fluorescent protein (GFP) reporter gene (Fig. The resultant gene construct, smyd1-gfp(5.3), was microinjected into zebrafish embryos for transient expression analysis. GFP expression in injected embryos was monitored by direct observation under a fluorescence microscope. Over 95% (n = 287) of the injected embryos exhibited GFP expression in skeletal muscles (Fig. In addition to the skeletal muscle expression, GFP expression was also observed in cranial, pectoral fin, and cardiac muscles (data not shown). However, very little expression was detected in nonmuscle tissues (data not shown).

These data indicated that the 5.3-kb smyd1 promoter and 5′-flanking sequence contained the essential regulatory sequence for muscle-specific expression. Genomic structure of zebrafish smyd1 gene and construction of smyd1-gfp gene construct.

Zebrafish smyd1 gene contains 10 exons. The partial DNA sequence of the zebrafish smyd1 promoter is shown. The transcriptional start site (AGACG) and the seven E-boxes (a–g) are indicated. Rabbit β-globin intron-2 sequence (in) and SV40 polyadenylation and transcription termination signals (sv) are included in the smyd1-gfp constructs to ensure proper processing of the RNA transcripts. The translation start site (ATG) is underlined. Transient expression analysis of smyd1-gfp in zebrafish embryos. A–F: GFP expression in skeletal muscles of zebrafish embryos injected with smyd1-gfp constructs that contain the 5.3 kb (A,B), 2 kb (C,D), or 0.5 kb (E,F) smyd1 gene promoter and 5′-flanking sequences, respectively.

Scale bars = 500 μm in A,C,E, 100 μm in B,D,F. Characterization of smyd1 Expression Using smyd1-gfp Transgenic Zebrafish To confirm the muscle specificity of the 5.3-kb smyd1 promoter and 5′-flanking sequence, and to determine whether the smyd1 is expressed in other striated muscles, we next generated transgenic zebrafish lines carrying the smyd1-gfp transgene. Five smyd1-gfp transgenic lines were generated. The temporal and spatial GFP expression was analyzed in these transgenic lines. GFP expression was first detected in adaxial cells flanking the notochord that give rise to slow muscles (Fig. As somites further develop and mature, GFP expression was identified in the presomitic mesoderm and developing somites (Fig.

GFP expression was also observed in the tail bud. However, the tail bud expression was restricted to presomitic mesoderm but not in the axial mesoderm (Fig. In addition to expression in the presomatic mesoderm and developing somite and skeletal muscles, GFP expression was also detected in cardiac, pectoral fin, and cranial muscles (Fig. The expression in skeletal and cardiac muscles is consistent with the expression pattern of smyd1 mRNA (Fig.

M; Tan et al., ). Expression of the smyd1-gfp transgene in the cranial and fin muscles was, however, a new finding for the smyd1 gene. To determine whether the endogenous smyd1 gene was expressed in cranial and fin muscles, in situ hybridization was performed with a smyd1 antisense probe in zebrafish embryos. The results showed that smyd1 mRNAs were indeed expressed in cranial and fin muscles in zebrafish embryos, although signals from in situ hybridization appeared weak in these tissues (Fig. These data demonstrated that, in addition to skeletal and cardiac muscles, smyd1 is also expressed in other striated muscles, including cranial muscles and pectoral fin muscles. To further confirm that the cranial GFP-expressing cells are indeed cranial muscles, we performed immunostaining with MF20 antibody (Bader et al., ), which labels cranial muscles in zebrafish (Hernandez et al., ). The pattern of MF20 staining appeared identical to GFP expression in smyd1-gfp transgenic fish (Fig.

Together, these data suggest that the 5.3-kb promoter and 5′-flanking sequence contained the necessary regulatory elements for smyd1 muscle-specific expression in cranial, cardiac, skeletal, and pectoral fin muscles. Expression of green fluorescent protein (GFP) reporter gene in developing somites, skeletal, cardiac, head, and fin muscles of transgenic embryos. A–C: In situ hybridization showing expression of smyd1 mRNA in 14 (A), 16 (B), and 22 (C) hours postfertilization (hpf) zebrafish embryos. D–F: GFP expression in developing somites and skeletal muscles of 14 (D), 16 (E), and 22 (F) hpf smyd1-gfp transgenic embryos. G,H: High magnification shows smyd1 mRNA (G) and smyd1-gfp (H) expression in presomitic mesoderm in the tail bud of 24 hpf embryos. I: Dorsal view of GFP expression in presomitic mesoderm but not in the axial mesoderm in the tail bud of smyd1-gfp transgenic embryos at 24 hpf.

The arrow indicates axial mesoderm. J: Side view of GFP expression in cardiac muscles of smyd1-gfp transgenic fish at 2 days postfertilization (dpf). K,L: GFP expression in pectoral fin (K, dorsal view) and cranial muscles (L, ventral view) at 2 or 3 dpf, respectively. M–O: In situ hybridization showing expression of smyd1 mRNA in cardiac (M), pectoral fin (N), and cranial muscles (O). Arrows in M and N indicate heart and pectoral fin, respectively.

P,Q: Expression of GFP (P, dorsal view) or smyd1 mRNA (Q, side view) in eye muscles of smyd1-gfp transgenic larvae at 5 dpf. R,S: Colocalization of GFP expression and MF20 staining in craniofacial muscles at 4 dpf.

GFP expression (R) and MF20 staining (S) were directly observed on the same smyd1-gfp transgenic embryo using a green or a red filter, respectively. Scale bars = 150μm in A–C,R,S, 30μm in G,H, 15 μm in I, 200 μm in M–O,P,Q. Characterization of a Key Regulatory Region in the smyd1 Promoter To identify the key regulatory region responsible for the muscle-specific expression, we performed a deletion analysis within the smyd1 5.3-kb promoter and its flanking sequence.

GFP constructs with different size smyd1 promoter and flanking sequences were generated and analyzed by transient expression in zebrafish embryos (Fig. Deleting the 5′-flanking region up to −0.5 kb upstream of the smyd1 promoter had little effect on the activity and tissue-specificity of the promoter (Fig. GFP expression remained strong and specific in skeletal, cranial, cardiac, and fin muscles (Fig.

These results indicated that the key cis-acting element(s) that regulates muscle-specific expression is located within the 0.5-kb smyd1 promoter region. Analyses of the 0.5-kb smyd1 promoter sequence identified seven putative E-box sites (CAnnTG) that might be involved in the muscle-specific expression (Fig.

Supporting this idea, several conserved E-box sites have also been found in the proximal promoter sequences in smyd1 genes of fugu, mouse, and human (Phan et al., ). To determine the role of these E-box sites in muscle-specific expression, the E-box sites in the smyd1 promoter were mutated individually or in various combinations.

The mutant constructs were injected into zebrafish embryos for transient expression analysis. Mutating any one of these seven E-boxes alone had little or no effect on the muscle specificity and activity of the smyd1 promoter. GFP expression from these single E-box mutations was comparable with the wild-type construct (data not shown).

However, mutating multiple E-box sites gradually reduced the GFP expression in skeletal muscles (Fig. Mutating all seven E-boxes resulted in significant reduction of GFP expression in skeletal muscles (Fig. It should be noted that GFP expression was not completely inhibited in skeletal muscles.

These data indicated that these E-box sites in the smyd1 promoter were involved in the muscle-specific expression, however, additional regulatory sequences might also be involved in regulating smyd1 gene expression in skeletal muscles. Characterization of E-box mutations on the activity of smyd1 promoter by transient expression analysis in zebrafish embryos. All panels are side views showing green fluorescent protein (GFP) expression in skeletal muscles of zebrafish embryos at 38 hours postfertilization (hpf). From A to H, zebrafish embryos were injected with the following DNA constructs: A: smyd1-gfp ( 0.5); B: smyd1-gfp ( a.); C: smyd1-gfp ( a. b.); D: smyd1-gfp ( a. b. c.); E: smyd1-gfp ( a.

b. c. d.); F: smyd1-gfp ( a. b. c. d.

e.), G: smyd1-gfp ( a. b. c. d. e. f.), and H: smyd1-gfp ( a.

b. c. d. e. f. g.), respectively.

Scale bar = 80 μm. Different Levels of GFP Expression in Slow and Fast Muscles of smyd1-gfp Transgenic Fish To further characterize GFP expression in smyd1-gfp transgenic fish, we analyzed GFP expression on cross-sections of skeletal muscles. Of interest, different levels of GFP expression were found in slow and fast muscles of smyd1-gfp transgenic fish. It appeared that slow muscles expressed higher levels of GFP than fast muscles (Fig. This finding was true for multiple transgenic zebrafish lines in both larval and adult stages (Fig. To test if smyd1 mRNA is expressed differently in slow and fast muscles, smyd1 mRNA expression was analyzed in zebrafish larvae by whole-mount in situ hybridization and followed by cross-section.

The results showed smyd1 expression in both slow and fast muscles of zebrafish larvae, although the staining signal appeared slightly stronger in superficial slow muscles compared with fast muscles (Fig. Different levels of green fluorescent protein (GFP) expression in slow and fast muscles of smyd1-gfp transgenic fish. A–C: GFP expression directly photographed on embryonic sections at 4 days postfertilization (dpf) under a confocal microscope. Slow muscles express higher levels of GFP than fast muscles in three transgenic lines of smyd1-gfp transgenic fish. A, line-27; B, line-32; C, line-51. Arrows indicate slow muscles. D,E: Cross-section showing higher levels of GFP expression in slow muscles of line-32 at 2 months old.

Slow muscles are indicated by arrows. F: Cross-section shows smyd1 mRNA expression in both slow and fast muscles at 4 dpf, although the staining appeared slightly stronger in superficial slow muscles. G,H: Cross-sections (dorsal on top) showing GFP expressing slow muscles in wild-type transgenic larvae (G) or yot mutant larvae (H) at 4 dpf. Slow muscles are clearly present at the dorsal and ventral myotome in yot mutant embryos. Scale bars = 150 μm in A–C, 500 μm in D, 200 μm in E, 120 μm in G,H. It has been shown that distinct mechanisms regulate slow muscle development at embryonic and larval stages (Barresi et al., ). Hh signaling is essential for the development of embryonic slow muscles, but not for larval slow muscles.

To determine the role of Hh signaling on smyd1-gfp expression, and to confirm that the stronger GFP expressing cells are indeed slow muscles, we analyzed GFP expression in yot mutant ( gli2 −/−) smyd1-gfp transgenic zebrafish larvae. The results showed that the gli2 mutation resulted in the absence of GFP-expressing slow muscles in the middle portion of the myotome. However, slow muscles were clearly present in the dorsal and ventral regions of the myotome (Fig. These data confirmed that the superficial cells expressing GFP in the middle region of the myotome are indeed slow muscles. Moreover, the results also reinforced the idea that development of slow muscles in the dorsal and ventral myotome is independent of Hh signaling at the larval stage.

To test if Hh signaling is involved in smyd1 expression in embryonic slow muscles at an early stage, we analyzed smyd1 mRNA expression in zebrafish mutants with Hh signaling defects. The results showed that smyd1 expression in adaxial cells was reduced in smu or yot mutant embryos that carry mutations in the Hedgehog receptor, Smoothened, or effector Gli2 (Fig. However, it should be noted that, although the smyd1 expression in adaxial cells was reduced, it was not completely diminished in these mutants. Two stripes of adaxial cells with weak smyd1 expression were clearly present in these mutants (Fig. E,G), suggesting that Hh was not required for initiation of smyd1 gene expression, but might be involved in maintaining its expression. Regulation of smyd1 gene expression in adaxial cells by Hedgehog signal. A–C: Microinjection of Shh mRNA induces smyd1 (B) or myod (C) expression in the injected side as indicated by arrows.

D–G: Comparison of smyd1 expression in wild-type (D,F) and smo (E) or yot (G) mutant embryos. Smyd1 expression is reduced in the mutant embryos. F and G are dual in situ with smyd1 (blue) and myod (red). Scale bars = 200 μm in A–C, 150 μm in D, 75 μm in E.

To confirm the importance of Hh signaling in smyd1 gene expression, we performed a reciprocal experiment by analyzing smyd1 expression in zebrafish embryos with ectopic Sonic Hh (Shh) expression. Overexpression of Shh by mRNA microinjection induced smyd1 expression in paraxial mesoderm in zebrafish embryos (Fig. Compared with the noninjected side, smyd1 mRNA expression was significantly expanded in the Shh mRNA-injected side (Fig. Together, these data indicate that Hh signal may be important for maintaining smyd1 gene expression in adaxial cells.

DISCUSSION In this study, we investigated smyd1 gene expression and regulation of its muscle-specific expression. We showed that smyd1 is specifically expressed in striated muscles, including skeletal, cardiac, cranial, and fin muscles. GFP expression in smyd1-gfp transgenic fish recapitulated that of the endogenous smyd1 gene. Hh signal appeared to be important for maintenance of smyd1 gene expression in slow muscles.

To the best of our knowledge, this is the first report of smyd1 gene expression in cranial muscles. This study provides new insights into the expression pattern of smyd1 gene, and moreover, several useful transgenic zebrafish models to study the molecular regulation of muscle cell differentiation. Regulation of smyd1 Muscle-Specific Expression We demonstrated that the 5.3-kb smyd1 gene promoter and its 5′-flanking sequence could drive GFP expression in a muscle-specific pattern in zebrafish embryos. The key regulatory sequence appeared to be located within the 0.5-kb proximal promoter region, because deletion of the 5′-flanking sequence up to 0.5 kb of the proxima l promoter had little or no effect on the tissue specificity and the activity of the promoter in skeletal muscles.

Because of the mosaic problem with the transient expression assay, the key regulatory sequences for smyd1 expression in cardiac, cranial, and fin muscles were not determined. Sequence analyses revealed that the proximal promoter contained seven putative E-box sites for binding of myogenic regulatory factors (MRFs). Several of these E-box sites are conserved in the smyd1 promoter sequences of other species, suggesting that they might be involved in regulation of smyd1 expression. Mutating these E-Box sites alone had little effect on the promoter activity. However, mutating seven E-boxes together significantly reduced the GFP expression of the smyd1-gfp constructs in skeletal muscles. These data argue that the E-box binding MRFs, such as MyoD, Myf-5, MRF-4, and Myogenin, may regulate smyd1 gene expression in skeletal muscles. Consistent with this idea, recent studies by Phan and colleagues (Phan et al., ) showed that three E-boxes within the −637-bp regulatory sequence region of smyd1 gene were required for its expression in developing skeletal muscles in mice.

further demonstrated that MyoD and Myogenin could bind directly to the promoter sequence of smyd1 and up-regulate smyd1 mRNA expression in transfected C2C12 myotubes. Although these studies indicate that MRFs may regulate smyd1 expression. It should be noted that mutating all seven E-boxes in the zebrafish smyd1 gene regulatory region did not completely abolish its activity in directing GFP expression in skeletal muscles, suggesting that other regulatory element(s) might be involved in its expression. Supporting this hypothesis, Blais and colleagues have showed that Mef2 could also bind to the promoter sequence of smyd1 gene and activate its expression in C2C12 cells.

However, Phan et al. demonstrated that the Mef2 site at −329/−320 of mouse smyd1 promoter was important for smyd1 expression in cardiac muscles, but not in skeletal muscles. Analyses of zebrafish smyd1 promoter sequence revealed that it contains several potential A/T-rich MEF2 binding sites in the proximal promoter region. However, their functions in controlling smyd1 expression remain to be determined. Regulation of muscle gene expression by multiple regulatory sequences appears to be a common mechanism for the control of muscle-specific gene expression.

Because smyd1 gene is expressed in several types of muscles, it remains to be determined if the same or distinct regulatory elements are responsible for its expression in skeletal, cardiac, cranial, and fin muscles using the transgenic approach. Regulation of smyd1 Expression and Adaxial Cell Formation by Hedgehog Signaling Hedgehog signals from the notochord and floor plate are required for differentiation of adaxial cells to slow muscles (Currie and Ingham,; Blagden et al.,; Du et al., ). Zebrafish mutants with a defective Hh signaling pathway, such as yot and smu that carry mutations in the Gli2 and Smoothened, respectively, lack embryonic slow muscles (Norris et al.,; Du and Dienhart,; Schauerte et al.,; Barresi et al., ). Questions have been raised regarding what happened to the adaxial cells in yot or smu mutants. We showed by in situ hybridization that the smyd1 gene was expressed in adaxial cells adjacent to the notochord in yot and smu mutant embryos, although, at a reduced level. These data indicate that Hh signal is not essential for smyd1 expression in adaxial cells.

Moreover, it also suggests that specification of adaxial cells does not require Hh signaling. These data are consistent with previous findings by Hirsinger and colleagues who showed that Hh signal is required for commitment but not initial induction of slow muscle precursors in zebrafish embryos (Hirsinger et al., ). Adaxial cells could form in the yot mutant, however, without Hh signaling, adaxial cells fail to differentiate into slow muscles, instead they differentiate into fast muscles. Distinct Patterns of smyd1 Expression in Slow and Fast Muscles and Hh Signaling in Development of Slow Muscles Analyses of multiple smyd1-gfp transgenic lines revealed distinct patterns of GFP expression in slow and fast muscles. It appears that slow muscles have a stronger GFP expression than fast muscles.

However, we could not state conclusively that smyd1 is preferably expressed in slow muscles, because in situ hybridization analyses showed that smyd1 mRNAs were expressed in both slow and fast muscles, although the staining appeared slightly stronger in superficial slow muscles. Therefore, it is not clear whether the higher levels of GFP expression observed in slow muscles should be considered as an ectopic event. Barresi and colleagues have demonstrated that, unlike embryonic slow muscles, formation of slow muscles at the larval stage did not require Hh signal. Yot or smu mutant embryos could still develop slow muscle fibers at the dorsal and ventral regions of the myotome, termed growth zones. We analyzed GFP expression in yot mutant smyd1-gfp transgenic fish at the larval stage.

Consistent with previous findings by Barresi and colleagues , we showed that GFP expression was missing in embryonic slow muscles, but slow muscle fibers expressing GFP were indeed present at the dorsal and ventral regions of the myotome in yot mutant transgenic fish at larval stage. Our findings reinforce the idea proposed by Barresi and colleagues that distinct mechanisms regulate slow muscle development at embryonic and larval stages. EXPERIMENTAL PROCEDURES Collection and Fertilization of Zebrafish Eggs and Raising of Zebrafish Embryos Mature zebrafish were raised at the zebrafish facility at the Aquaculture Research Center, University of Maryland Biotechnology Institute. The fish were maintained at a photoperiod of 14 hr light and 10 hr dark in 8-gallon aquariums supplied with freshwater and aeration.

Spawning of zebrafish was carried out by putting one male in the tank with two females. The mutant strains used in this study were smu mutant allele ( smu b641) and yot mutant allele ( yot ty119; van Eeden et al.,; Varga et al., ). Determination of Transcription Start Site by 5′-RACE RACE was carried out to determine the transcription start site according to the instructions provided by SMART RACE cDNA Amplification Kit (BD Bioscience, Clonetech Lab., Inc). The 5′-RACE-Ready cDNA was amplified with the adapter primer (see manual protocol) and a 5′ gene-specific primer (5′-agagtcgtcagctgattgtcg-3′). The polymerase chain reaction (PCR) product was subcloned into the pGEM-T easy vector for sequencing.

DNA Constructs The smyd1-gfp construct was made by linking the 5.3-kb smyd1 promoter and its flanking sequence with the GFP reporter gene. The 5.3-kb sequence of smyd1 was identified by blasting zebrafish genome sequence in GenBank. The sequence was isolated by PCR using two gene-specific primers (smyd1-5′-UTR and smyd1-5′-flank) based on the DNA sequences in the 5′-UTR and upstream 5′ flanking sequence, respectively.

SalI sites were introduced at both ends of the 5.3-kb sequence by the PCR primers for subcloning. The 5.3-kb DNA sequence was first cloned into the T-easy vector (Promega).

The sequence was then released from the vector by SalI digestion and subcloned into the SalI site of a GFP vector (IGFP) that contained a 640-bp rabbit β- globin intron 2 sequence in front of the GFP reporter gene (Du and Dienhart, ). The rabbit β- globin intron 2 sequence has been shown to enhance gene expression in transfected culture cells and in transgenic mice and transgenic fish (Buchman and Berg,; Brinster et al.,; Amsterdam et al.,; Du and Dienhart, ). To facilitate the linearization by SalI digestion, the second SalI site at the junction between the promoter and the rabbit β- globin intron sequence was subsequently mutated by partial SalI digestion and followed by Klenow filling-in and self-regulation. The final construct was named smyd1-gfp(5.3). To generate constructs with different size promoter sequences, PCR was performed using the 5′-UTR primer together with specific primers, smyd1-4k, smyd1-3k, smyd1-2k, smyd1-1k, or smyd1-0.5k, which are located at approximately 4, 3, 2, 1, and 0.5 kb from the ATG start site, respectively. The resultant PCR products were cloned upstream of the IGFP vector at the SalI site to produce smyd1-gfp (4), smyd1-gfp (3), smyd1-gfp (2), smyd1-gfp (1), and smyd1-gfp (0.5) constructs that contained the 4 kb, 3 kb, 2 kb, 1 kb, and 0.5 kb of the smyd1 promoter and 5′-flanking sequences, respectively.

The seven E-box consensus sequences CAxxTG; from a to g in the smyd1-gfp (0.5) construct were mutated to C Gxx CG individually or in combinations using the QuickChange site-directed mutagenesis kit (Stratagene). The resulting plasmids with single E-box mutations were named smyd1-gfp ( a.), smyd1-gfp ( b.), smyd1-gfp ( c.), smyd1-gfp ( d.), smyd1-gfp ( e.), smyd1-gfp ( f.), and smyd1-gfp ( g.), respectively. The resulting plasmids with multiple mutations were named smyd1-gfp ( a. b.), smyd1-gfp ( a. b. c.), smyd1-gfp ( a.b.c.d.), smyd1-gfp ( a.

b. c. d.

e.), smyd1-gfp ( a. b. c. d. e. f.), and smyd1-gfp ( a.

b. c. d. e.

f. g.), respectively. The sequences and positions of these E-boxes were mutated as follows: E-box a ( −305CAGCTG −300 to C GGC CG); E-box b ( −277CAACTG −272 to C GAC CG); E-box c ( −245CACATG −240 to C GCA CG); E-box d ( −221CACATG −216 to C GCA CG); E-box e ( −174CAGCTG −169 to C GGC CG); E-box f ( −142CATTTG −137 to C GTT CG), and E-box g ( −47CAGCTG −42 to C GGC CG). Primers used were as follows: smyd1-5′flank, 5′-GTCGACGTTCACAGGACTCCTTAGGGTTAG; smyd1-4k, 5′-CACTGTTGCGACAACATTA; smyd1-3k, 5′-GTAAGCAGAAATGTGTGTAA; smyd1-2k, 5′-TGAGAGTCTGATCTAGACTGT; smyd1-1k, 5′-GACGTGAGATTGTGTTGTCT; smyd1-0.5k, 5′-GTCGACGATCAGATCTGGTATGCAGTAGT; and smyd1-5′-UTR, 5′-GTCGACCTGGATCTTCAGCGTCTGCA. Microinjection and Transient Expression Analyses in Zebrafish Embryos smyd1-gfp DNA constructs that contained various sizes of the smyd1 5′-flanking sequence or E-box mutations were linearized at the 5′ end by SalI digestion.

The linearized DNA plasmid was purified using a Gel Extraction kit (Qiagen). The resultant DNA plasmid was dissolved in distilled H 2O to a final concentration of 50 μg/ml. Approximately 2 nl of DNA solution was microinjected into the cytoplasm of zebrafish embryos at the one- or two-cell stage. Microinjection was carried out under a dissection microscope (MZ8, Leica) using a PLI-100 pico-injector (Medical System Corp.).

GFP expression in the injected zebrafish embryos was analyzed at 24–48 hours postfertilization (hpf). GFP expression was analyzed by direct observation of GFP expression under a fluorescent microscope or using anti-GFP antibody staining.

The numbers of embryos showed GFP expression, and the numbers of fibers expressing GFP were determined and compared among different gene constructs to score the activity and tissue-specificity of these promoter sequences. Production and Screening of smyd1-gfp(5.3) Transgenic Zebrafish Transgenic zebrafish were generated for the smyd1-gfp(5.3) DNA construct.

Linearized DNA plasmid ( SalI cut) was injected into fertilized zebrafish eggs as described above. Germline transgenic founders were identified by screening their F1 embryos for GFP expression under a fluorescence microscope. Five transgenic fish (#14, #23, #27, #32, and #51) carrying the smyd1-gfp(5.3) DNA construct were identified. Approximately 100–200 F1 embryos of these transgenic founders were allowed to grow to maturity. Adult transgenic fish of the F1 generation were screened individually by PCR using DNA extracted from their caudal fins. To clip fins, fish were anesthetized in 0.02% MS-222.

A small piece of tissue from the caudal fin was cut and collected in a 1.5-ml microfuge tube. Tissue of the caudal fin was added with 200 μl of lysis buffer containing 50 mM KCl, 10 mM Tris pH 8.8, 1.5 mM MgCl 2, 0.1% Triton X-100. The sample was boiled for 5 min and was then digested in proteinase K (100 μg/ml) for 1 hr at 55°C. Proteinase K was inactivated by boiling for 5 min after the digestion. The samples were centrifuged at 12 k for 5 min, and 1 μl of the supernatant was used for each PCR reaction. Smyd1-gfp transgene-specific primers (smyd1-5′-5 and GFP-R) were used for PCR analysis. Smyd1-5′-5 was derived from the 5′-flanking sequence of smyd1, while GFP-R was derived from the antisense strand of GFP coding sequence.

Control PCR primers (HH-F and HH-R) were derived from the sense and antisense strand of tiggy-winkle hedgehog exon 3 sequence, respectively. PCR using this set of primers produced a 615-bp DNA fragment. Smyd1-5′-5, 5′-CTGACATGGTACAGCTGAAGA-3′; GFP-R, 5′-GCCATGTGTAATCCCAGCAGC-3′; HH-F, 5′-GGACGGTGACACTTGGTGATG-3′; HH-R, 5′-CGAGTGGATGGAAAGAGTCTC-3′ PCR was carried out in a 25-μl reaction solution containing 1 μM of each primer, four deoxyribonucleotide triphosphate at 0.2 mM for each, and 0.5 units of Taq DNA polymerase (Promega).

PCR was carried out for 35 cycles. Each cycle included 30 sec at 94°C, 30 sec at 58°C, and 1 min at 72°C. Ten microliters of the amplified product was analyzed by electrophoresis on a 1% agarose gel. Whole-Mount Antibody Staining With MF20 Antibody Whole-mount antibody staining was performed on smyd1-gfp transgenic larvae at 4 days postfertilization (dpf) using MF20 antibody (Hybridoma Bank). Zebrafish larvae were fixed in 4% paraformaldehyde in 1× phosphate buffered saline (PBS) for 1 hr at room temperature, and then washed with PBST (1× PBS, 0.1% Tween) for 3× 10 min.

The larvae were then incubated with collagenase (1 mg/ml; Sigma C-9891) for 2 hr at room temperature. The larvae were washed with PBST for 2× 10 min and then treated with cold acetone (−20°C) for 10 min. For whole-mount immunostaining, the larvae were washed with PBST for 2× 10 min and then blocked with 10% goat serum for 60 min at room temperature. The larvae were subsequently incubated with 1:20 diluted anti-myosin MF20 antibody overnight at 4°C. The larvae were then washed 3× 20 min with PBST, followed by incubation with 1:500 diluted TRITC-conjugated secondary antibody (Sigma T-7657) in PBST for 1 hr at room temperature.

The larvae were then washed 3× 20 min in PBST and covered with VECTASHIELD (Vector Laboratories). The larvae were photographed under a fluorescent microscopy (Axioplan-2, Zeiss) using a green or a red filter. Cryostat Sectioning and Direct GFP Observation Embryos of 4 dpf or adult fish were fixed in 4% paraformaldehyde in 1×PBS for 1 hr at room temperature, and then washed with PBS 3×10 min. Next the fixed embryos and fish were soaked in 30% sucrose until they sank. The fixed samples were then transferred into an embedding chamber filled with OCT cryostat embedding medium (Tissue Tek). The embedding chamber was frozen on dry ice.

Axion Dpf 8505pt

Frozen blocks were cut on a cryostat at −20°C to produce 10-μm sections. Sections were transferred to subbed slides and allowed to dry completely. For direct observation of GFP expression, the sections were rehydrated in PBS-Tween (PBS, 0.1% Tween), covered with VECTASHIELD (H-1000, Vector Laboratories), and photographed under a confocal microscope directly.

One-Color and Two-Color Whole-Mount In Situ Hybridization The plasmid clone ( pGEM-zsmyd1) containing the smyd1 cDNA sequence of exon 9 and 10 and the 3′-UTR region was used as a template for synthesizing the antisense digoxigenin-labeled RNA probe. The plasmid was linearized with EcoRI and transcribed with T7 RNA polymerase. For double in situ hybridization, zebrafish MyoD antisense probe was labeled with fluorescein. In situ hybridization with a single probe (one color) was carried out as described by Du and Dienhart.

Two-color double in situ hybridization was carried out as described by Tan and Du using the fluorescein-labeled MyoD probe and the digoxigenin-labeled smyd1 probe. Embryos were photographed in 3% methylcellulose.

Cyclopamine Treatment of Zebrafish Embryos Cyclopamine treatment was carried out as described by Barresi et al. Briefly, cyclopamine (Toronto Research Chemical, Toronto, CA) was dissolved in dimethyl sulfoxide to make a 10 mM stock solution. Embryos were treated with cyclopamine at 100 μM in 35-mm dishes. Cyclopamine was added at shield stage, and it was left in until fixation for in situ hybridization.

Acknowledgements Special thanks to Steve Devoto from Wesleyan University for providing zebrafish mutant strains. The MF20 monoclonal antibody developed by D.A.

Fischman was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by The University of Iowa, Department of Biological Sciences (Iowa City, IA 52242). Josep Rotllant is a Fulbright scholar supported by a Fulbright fellowship from Spain. This publication is contribution number 06-146 from the Center of Marine Biotechnology, University of Maryland Biotechnology Institute.

Ancillary Article Information. Amsterdam A, Lin S, Hopkins N.

The Aequorea victoria green fluorescent protein can be used as a reporter in live zebrafish embryos. Dev Biol 171: 123– 129. Bader D, Masaki T, Fischman DA. Immunochemical analysis of myosin heavy chain during avian myogenesis in vivo and in vitro. J Cell Biol 95: 763– 770.

Barresi MJ, Stickney HL, Devoto SH. The zebrafish slow-muscle-omitted gene product is required for Hedgehog signal transduction and the development of slow muscle identity. Development 127: 2189– 2199.

Barresi MJ, D'Angelo JA, Hernandez LP, Devoto SH. Distinct mechanisms regulate slow-muscle development. Curr Biol 11: 1432– 1438. Biron VL, McManus KJ, Hu N, Hendzel MJ, Underhill DA. Distinct dynamics and distribution of histone methyl-lysine derivatives in mouse development. Dev Biol 276: 337– 351.

Blagden CS, Currie PD, Ingham PW, Hughes SM. Notochord induction of zebrafish slow muscle mediated by Sonic hedgehog. Genes Dev 11: 2163– 2175. Blais A, Tsikitis M, Acosta-Alvear D, Sharan R, Kluger Y, Dynlacht BD. An initial blueprint for myogenic differentiation. Genes Dev 19: 553– 569.

Brinster R.J, Behringer AR, Gelia P, Palmiter R. Intron increase transcriptional efficiency in transgenic mice. Proc Natl Acad Sci U S A 85: 836– 840. Buchman A, Berg P. Comparison of intron-dependent and intron-independent gene expression. Mol Cell Biol 8: 4395– 4405.

Cerny J, Quesenberry PJ. Chromatin remodeling and stem cell theory of relativity. J Cell Physiol 201: 1– 16. Cheng X, Collins RE, Zhang X. Structural and sequence motifs of protein (histone) methylation enzymes.

Annu Rev Biophys Biomol Struct 34: 267– 294. Cirillo L, Zaret K. Developmental biology. A linker histone restricts muscle development.

Science 304: 1607– 1609. Currie PD, Ingham PW. Induction of a specific muscle cell type by a Hedgehog-like protein in zebrafish. Nature 382: 452– 455.

Du SJ, Dienhart M. Gli2 mediation of hedgehog signals in slow muscle induction in zebrafish. Differentiation 67: 84– 91. Du SJ, Dienhart M. Zebrafish tiggy-winkle hedgehog promoter directs notochord and floor plate green fluorescence protein expression in transgenic zebrafish embryos. Dev Dyn 222: 655– 666.

Du SJ, Devoto SH, Westerfield M, Moon RT. Positive and negative regulation of muscle cell identity by members of the Hedgehog and TGF-β gene families. J Cell Biol 139: 145– 156. Du SJ, Gao J, Anyangwe V. Muscle-specific expression of myogenin in zebrafish embryos is controlled by multiple regulatory elements in the promoter.

Comp Biochem Physiol B Biochem Mol Biol 134: 123– 134. Gelmetti V, Zhang J, Fanelli M, Minucci S, Pelicci PG, Lazar MA. Aberrant recruitment of the nuclear receptor corepressor-histone deacetylase complex by the acute myeloid leukemia fusion partner ETO. Mol Cell Biol 18: 7185– 191. Gottlieb PD, Pierce SA, Sims RJ, Yamagishi H, Weihe EK, Harriss JV, Maika SD, Kuziel WA, King HL, Olson EN, Nakagawa O, Srivastava D. Bop encodes a muscle-restricted protein containing MYND and SET domains and is essential for cardiac differentiation and morphogenesis.

Nat Genet 31: 25– 32. Hamamoto R, Furukawa Y, Morita M, Iimura Y, Silva FP, Li M, Yagyu R, Nakamura Y. SMYD13 encodes a histone methyltransferase involved in the proliferation of cancer cells. Nat Cell Biol 6: 731– 740. Hamamoto R, Silva FP, Tsuge M, Nishidate T, Katagiri T, Nakamura Y, Furukawa Y.

Enhanced SMYD3 expression is essential for the growth of breast cancer cells. Cancer Sci 97: 113– 118. Hernandez LP, Patterson SE, Devoto SH. The development of muscle fiber type identity in zebrafish cranial muscles. Anat Embryol (Berl) 209: 323– 334. Hirsinger E, Stellabotte F, Devoto SH, Westerfield M. Hedgehog signaling is required for commitment but not initial induction of slow muscle precursors.

Dev Biol 275: 143– 157. Hwang I, Gottlieb PD. Bop: a new T-cell-restricted gene located upstream of and opposite to mouse CD8b. Immunogenetics 42: 353– 361. Hwang I, Gottlieb PD. The Bop gene adjacent to the mouse CD8b gene encodes distinct zinc-finger proteins expressed in CTLs and in muscle. J Immunol 158: 1165– 1174.

Kubicek S, Jenuwein T. A crack in histone lysine methylation. Cell 119: 903– 906. Lee H, Habas R, Abate-Shen C.

MSX1 cooperates with histone H1b for inhibition of transcription and myogenesis. Science 304: 1675– 1678. Lutterbach B, Westendorf JJ, Linggi B, Patten A, Moniwa M, Davie JR, Huynh KD, Bardwell VJ, Lavinsky RM, Rosenfeld MG, Glass C, Seto E, Hiebert SW.

ETO, a target of t(8;21) in acute leukemia, interacts with the N-CoR and mSin3 corepressors. Mol Cell Biol 18: 7176– 7184. McKinsey TA, Zhang CL, Olson EN. Control of muscle development by dueling HATs and HDACs.

Curr Opin Genet Dev 11: 497– 504. McKinsey TA, Zhang CL, Olson EN.

Abdominal Manual Muscle Testing

Signaling chromatin to make muscle. Curr Opin Cell Biol 14: 763– 772. Norris W, Neyt C, Ingham PW, Currie PD. Slow muscle induction by Hedgehog signalling in vitro.

J Cell Sci 113: 2695– 2703. Phan D, Rasmussen TL, Nakagawa O, McAnally J, Gottlieb PD, Tucker PW, Richardson JA, Bassel-Duby R, Olson EN. BOP, a regulator of right ventricular heart development, is a direct transcriptional target of MEF2C in the developing heart. Development 132: 2669– 2678.

Rea S, Eisenhaber F, O'Carroll D, Strahl BD, Sun ZW, Schmid M, Opravil S, Mechtler K, Ponting CP, Allis CD, Jenuwein T. Regulation of chromatin structure by site-specific histone H3 methyltransferases. Nature 406: 593– 599. Rupp RA, Singhal N, Veenstra GJ.

When the embryonic genome flexes its muscles. Eur J Biochem 269: 2294– 2299. Santos-Rosa H, Schneider R, Bannister AJ, Sherriff J, Bernstein BE, Emre NC, Schreiber SL, Mellor J, Kouzarides T. Active genes are tri-methylated at K4 of histone H3. Nature 419: 407– 411.

Schauerte HE, van Eeden FJ, Fricke C, Odenthal J, Strahle U, Haffter P. Sonic hedgehog is not required for the induction of medial floor plate cells in the zebrafish. Development 125: 2983– 2993. Sims RJ III, Reinberg D. From chromatin to cancer: a new histone lysine methyltransferase enters the mix.

Nat Cell Biol 6: 685– 687. Tan X, Du J. Differential expression of two MyoD genes in fast and slow muscles of gilthead seabream (Sparus aurata). Dev Genes Evol 212: 207– 217. Tan XG, Rotllant J, Li H, DeDeyne P, Du SJ. SmyD1, a histone methyltransferase, is required for myofibril organization and muscle contraction in zebrafish embryos.

Proc Natl Acad Sci U S A 103: 2713– 2718. van Eeden FJ, Granato M, Schach U, Brand M, Furutani-Seiki M, Haffter P, Hammerschmidt M, Heisenberg CP, Jiang YJ, Kane DA, Kelsh RN, Mullins MC, Odenthal J, Warga RM, Allende ML, Weinberg ES, Nusslein-Volhard C. Mutations affecting somite formation and patterning in the zebrafish, Danio rerio. Development 123: 153– 164. Varga ZM, Amores A, Lewis KE, Yan YL, Postlethwait JH, Eisen JS, Westerfield M. Zebrafish smoothened functions in ventral neural tube specification and axon tract formation.

Development 128: 3497– 3509. Wang J, Hoshino T, Redner RL, Kajigaya S, Liu JM. ETO, fusion partner in t(8;21) acute myeloid leukemia, represses transcription by interaction with the human N-CoR/mSin3/HDAC1 complex. Proc Natl Acad Sci U S A 95: 10860– 10865. Zhang CL, McKinsey TA, Olson EN. Association of class II histone deacetylases with heterochromatin protein 1: potential role for histone methylation in control of muscle differentiation. Mol Cell Biol 22: 7302– 7312.

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