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Homologous recombination provides high-fidelity DNA repair throughout all domains of life. Live cell fluorescence microscopy offers the opportunity to image individual recombination events in real time providing insight into the in vivo biochemistry of the involved proteins and DNA molecules as well as the cellular organization of the process of homologous recombination. Herein we review the cell biological aspects of mitotic homologous recombination with a focus on Saccharomyces cerevisiae and mammalian cells, but will also draw on findings from other experimental systems. Key topics of this review include the stoichiometry and dynamics of recombination complexes in vivo, the choreography of assembly and disassembly of recombination proteins at sites of DNA damage, the mobilization of damaged DNA during homology search, and the functional compartmentalization of the nucleus with respect to capacity of homologous recombination.Homologous recombination (HR) is defined as the homology-directed exchange of genetic information between two DNA molecules (Fig. 1). Mitotic recombination is often initiated by single-stranded DNA (ssDNA), which can arise by several avenues (Mehta and Haber 2014). They include the processing of DNA double-strand breaks by 5′ to 3′ resection, during replication of damaged DNA, or during excision repair (Symington 2014). The ssDNA is bound by replication protein A (RPA) to control its accessibility to the Rad51 recombinase (Sung 1994, 1997a; Sugiyama et al. 1997; Morrical 2014). The barrier to Rad51-catalyzed recombination imposed by RPA can be overcome by a number of mediators, such as BRCA2 and Rad52, which serve to replace RPA with Rad51 on ssDNA, and the Rad51 paralogs Rad55-Rad57 (RAD51B-RAD51C-XRCC2-XRCC3) and the Psy3-Csm2-Shu1-Shu2 complex (SHU) (RAD51D-XRCC2-SWS1), which stabilize Rad51 filaments on ssDNA (see Sung 1997b; Sigurdsson et al. 2001; Martin et al. 2006; Bernstein et al. 2011; Liu et al. 2011; Qing et al. 2011; Amunugama et al. 2013; Zelensky et al. 2014). The Rad51 nucleoprotein filament catalyzes the invasion into a homologous duplex to produce a displacement loop (D-loop) (Fig. 1). At this stage, additional antirecombination functions are exerted by Srs2 (FBH1, PARI), which dissociates Rad51 filaments from ssDNA, and Mph1 (FANCM), which disassembles D-loops (see Daley et al. 2014). Upon Rad51-catalyzed strand invasion, the ATP-dependent DNA translocase Rad54 enables the invading 3′ end to be extended by DNA polymerases to copy genetic information from the intact duplex (Li and Heyer 2009). Ligation of the products often leads to joint molecules (JMs), such as single- or double-Holliday junctions (s/dHJs) or hemicatenanes (HCs), which must be processed to allow separation of the sister chromatids during mitosis. JMs can be dissolved by the Sgs1-Top3-Rmi1 complex (STR) (BTR, BLM-TOP3α-RMI1-RMI2) (see Bizard and Hickson 2014) or resolved by structure-selective nucleases, such as Mus81-Mms4 (MUS81-EME1), Slx1-Slx4, and Yen1 (GEN1) (see Wyatt and West 2014). Mitotic cells favor recombination events that lead to noncrossover events likely to avoid potentially detrimental consequences of loss of heterozygosity and translocations.Open in a separate windowFigure 1.Primary pathways for homology-dependent double-strand break (DSB) repair. Recombinational repair of a DSB is initiated by 5′ to 3′ resection of the DNA end(s). The resulting 3′ single-stranded end(s) invade an intact homologous duplex (in red) to prime DNA synthesis. For DSBs that are repaired by the classical double-strand break repair (DSBR) model, the displaced strand from the donor duplex pairs with the 3′ single-stranded DNA (ssDNA) tail at the other side of the break, which primes a second round of DNA synthesis. After ligation of the newly synthesized DNA to the resected 5′ strands, a double-Holliday junction (dHJ) intermediate is generated. The dHJ can be either dissolved by branch migration (indicated by arrows) into a hemicatenane (HC) leading to noncrossover (NCO) products or resolved by endonucleolytic cleavage (indicated by triangles) to produce NCO (positions 1, 2, 3, and 4) or CO (positions 1, 2, 5, and 6) products. Alternatively to the double-strand break repair (DSBR) pathway, the invading strand is often displaced after limited synthesis and the nascent complementary strand anneals with the 3′ single-stranded tail of the other end of the DSB. After fill-in synthesis and ligation, this pathway generates NCO products and is referred to as synthesis-dependent strand annealing (SDSA).
Open in a separate windowThe vast majority of cell biological studies of mitotic recombination in living cells are performed by tagging of proteins with genetically encoded green fluorescent protein (GFP) or similar molecules (Shaner et al. 2005; Silva et al. 2012). In this context, it is important to keep in mind that an estimated 13% of yeast proteins are functionally compromised by GFP tagging (Huh et al. 2003). By choosing fluorophores with specific photochemical properties, it has been possible to infer biochemical properties, such as diffusion rates, protein–protein interactions, protein turnover, and stoichiometry of protein complexes at the single-cell level. To visualize the location of specific loci within the nucleus, sequence-specific DNA-binding proteins such the Lac and Tet repressors have been used with great success. Specifically, tandem arrays of 100–300 copies of repressor binding sites are inserted within 10–20 kb of the locus of interest in cells expressing the GFP-tagged repressor (Straight et al. 1996; Michaelis et al. 1997). In wild-type budding yeast, such protein-bound arrays are overcome by the replication fork without a cell-cycle delay or checkpoint activation (Dubarry et al. 2011). However, the arrays are unstable in rrm3Δ and other mutants (Dubarry et al. 2011). More pronounced DNA replication blockage by artificial protein-bound DNA tandem arrays has be observed in fission yeast, which is accompanied by increased recombination and formation of DNA anaphase bridges (Sofueva et al. 2011). Likewise, an array of Lac repressor binding sites was reported to induce chromosomal fragility in mouse cells (Jacome and Fernandez-Capetillo 2011). However, these repressor-bound arrays generally appear as a focus with a size smaller than the diffraction limit of light, which is in the range 150–300 nm for wide-field light microscopy. 相似文献
Table 1.
Evolutionary conservation of homologous recombination proteins between Saccharomyces cerevisiae and Homo sapiensFunctional class | S. cerevisiae | H. sapiens |
---|---|---|
End resection | Mre11-Rad50-Xrs2 | MRE11-RAD50-NBS1 |
Sae2 | CtIP | |
Exo1 | EXO1 | |
Dna2-Sgs1-Top3-Rmi1 | DNA2-BLM-TOP3α-RMI1-RMI2 | |
Adaptors | Rad9 | 53BP1, MDC1 |
– | BRCA1 | |
Checkpoint signaling | Tel1 | ATM |
Mec1-Ddc2 | ATR-ATRIP | |
Rad53 | CHK2 | |
Rad24-RFC | RAD17-RFC | |
Ddc1-Mec3-Rad17 | RAD9-HUS1-RAD1 | |
Dpb11 | TOPBP1 | |
Single-stranded DNA binding | Rfa1-Rfa2-Rfa3 | RPA1-RPA2-RPA3 |
Single-strand annealing | Rad52 | RAD52 |
Rad59 | – | |
Mediators | – | BRCA2-PALB2 |
Rad52 | – | |
Strand exchange | Rad51 | RAD51 |
Rad54 | RAD54A, RAD54B | |
Rdh54 | – | |
Rad51 paralogs | Rad55-Rad57 | RAD51B-RAD51C-RAD51D-XRCC2-XRCC3 |
Psy3-Csm2-Shu1-Shu2 | RAD51D-XRCC2-SWS1 | |
Antirecombinases | Srs2 | FBH1, PARI |
Mph1 | FANCM | |
– | RTEL | |
Resolvases and nucleases | Mus81-Mms4 | MUS81-EME1 |
Slx1-Slx4 | SLX1-SLX4 | |
Yen1 | GEN1 | |
Rad1-Rad10 | XPF-ERCC1 | |
Dissolution | Sgs1-Top3-Rmi1 | BLM-TOP3α-RMI1-RMI2 |
3.
Recent advances in the characterization of the archaeal DNA replication system together with comparative genomic analysis have led to the identification of several previously uncharacterized archaeal proteins involved in replication and currently reveal a nearly complete correspondence between the components of the archaeal and eukaryotic replication machineries. It can be inferred that the archaeal ancestor of eukaryotes and even the last common ancestor of all extant archaea possessed replication machineries that were comparable in complexity to the eukaryotic replication system. The eukaryotic replication system encompasses multiple paralogs of ancestral components such that heteromeric complexes in eukaryotes replace archaeal homomeric complexes, apparently along with subfunctionalization of the eukaryotic complex subunits. In the archaea, parallel, lineage-specific duplications of many genes encoding replication machinery components are detectable as well; most of these archaeal paralogs remain to be functionally characterized. The archaeal replication system shows remarkable plasticity whereby even some essential components such as DNA polymerase and single-stranded DNA-binding protein are displaced by unrelated proteins with analogous activities in some lineages.Double-stranded DNA is the molecule that carries genetic information in all cellular life-forms; thus, replication of this genetic material is a fundamental physiological process that requires high accuracy and efficiency (Kornberg and Baker 2005). The general mechanism and principles of DNA replication are common in all three domains of life—archaea, bacteria, and eukaryotes—and include recognition of defined origins, melting DNA with the aid of dedicated helicases, RNA priming by the dedicated primase, recruitment of DNA polymerases and processivity factors, replication fork formation, and simultaneous replication of leading and lagging strands, the latter via Okazaki fragments (Kornberg and Baker 2005; Barry and Bell 2006; Hamdan and Richardson 2009; Hamdan and van Oijen 2010). Thus, it was a major surprise when it became clear that the protein machineries responsible for this complex process are drastically different, especially in bacteria compared with archaea and eukarya. The core components of the bacterial replication systems, such as DNA polymerase, primase, and replication helicase, are unrelated or only distantly related to their counterparts in the archaeal/eukaryotic replication apparatus (Edgell 1997; Leipe et al. 1999).The existence of two distinct molecular machines for genome replication has raised obvious questions on the nature of the replication system in the last universal common ancestor (LUCA) of all extant cellular life-forms, and three groups of hypotheses have been proposed (Leipe et al. 1999; Forterre 2002; Koonin 2005, 2006, 2009; Glansdorff 2008; McGeoch and Bell 2008): (1) The replication systems in Bacteria and in the archaeo–eukaryotic lineage originated independently from an RNA-genome LUCA or from a noncellular ancestral state that encompassed a mix of genetic elements with diverse replication strategies and molecular machineries. (2) The LUCA was a typical cellular life-form that possessed either the archaeal or the bacterial replication apparatus in which several key components have been replaced in the other major cellular lineage. (3) The LUCA was a complex cellular life-form that possessed both replication systems, so that the differentiation of the bacterial and the archaeo–eukaryotic replication machineries occurred as a result of genome streamlining in both lines of descent that was accompanied by differential loss of components. With regard to the possible substitution of replication systems, a plausible mechanism could be replicon takeover (Forterre 2006; McGeoch and Bell 2008). Under the replicon takeover hypothesis, mobile elements introduce into cells a new replication system or its components, which can displace the original replication system through one or several instances of integration of the given element into the host genome accompanied by inactivation of the host replication genes and/or origins of replication. This scenario is compatible with the experimental results showing that DNA replication DNA in Escherichia coli with an inactivated DnaAgene or origin of replication can be rescued by the replication apparatus of R1 or F1 plasmids integrated into the bacterial chromosome (Bernander et al. 1991; Koppes 1992). Furthermore, genome analysis suggests frequent replicon fusion in archaea and bacteria (McGeoch and Bell 2008); in particular, such events are implied by the observation that in archaeal genomes, genes encoding multiple paralogs of the replication helicase MCM and origins of replication are associated with mobile elements (Robinson and Bell 2007; Krupovic et al. 2010). Replicon fusion also is a plausible path from a single origin of replication that is typical of bacteria to multiple origins present in archaea and eukaryotes. However, all the evidence in support of frequent replicon fusion and the plausibility of replicon takeover notwithstanding, there is no evidence of displacement of the bacterial replication apparatus with the archaeal version introduced by mobile elements, or vice versa, displacement of the archaeal machinery with the bacterial version, despite the rapid accumulation of diverse bacterial and archaeal genome sequences. Thus, the displacement scenarios of DNA replication machinery evolution are so far not supported by comparative genomic data.Regardless of the nature of the DNA replication system (if any) in the LUCA and the underlying causes of the archaeo–bacterial dichotomy of replication machineries, the similarity between the archaeal and eukaryotic replication systems is striking (Leipe et al. 1999; Bell and Dutta 2002; Bohlke et al. 2002; Kelman and White 2005; Barry and Bell 2006). Thus, the archaeal replication system appears to be an ancestral version of the eukaryotic system and hence a good model for functional and structural studies aimed at gaining mechanistic insights into eukaryotic replication.
Open in a separate windowFor eukaryotic genes in Homo sapiens and Saccharomyces cerevisiae, gene names are indicated. Archaeal genes are denoted as in Barry and Bell (2006) or as introduced here.aNot confidently traced to LACA.In the last few years, there has been substantial progress in the study of the archaeal replication systems that has led to an apparently complete delineation of all proteins that are essential for replication (Berquist et al. 2007; Beattie and Bell 2011a; MacNeill 2011). The combination of experimental, structural, and bioinformatics studies has led to the discovery of archaeal homologs (orthologs) for several components of the replication system that have been previously deemed specific for eukaryotes (Barry and Bell 2006; MacNeill 2010, 2011; Makarova et al. 2012). Furthermore, complex evolutionary events that involve multiple lineage-specific duplications, domain rearrangements, and gene loss, and in part seem to parallel the evolution of the evolution of the replication system in eukaryotes, have been delineated for a variety of replication proteins in several archaeal lineages (Tahirov et al. 2009; Chia et al. 2010; Krupovic et al. 2010). Here we summarize these findings and present several additional case studies that show the complexity of evolutionary scenarios for the components of the archaeal replication machinery and new aspects of their relationship with the eukaryotic replication system. 相似文献
Table 1.
The relationship between archaeal and eukaryotic replication systemsArchaea (projection for LACA) | Eukaryotes (projection for LECA) | Comments |
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ORC complex | ||
arORC1 | Orc1, Cdc6 | In LACA the ORC/Cdc6 complex probably consisted of two distinct subunits, and in LECA of six distinct. Both complexes might possess additional Orc6 and Cdt1 components. |
arORC2 | Orc2, Orc3, Orc4, Orc5 | |
TFIIB or homologa | Orc6 | |
WhiP or other wHTH proteina | Cdt1 | |
CMG complex | ||
Archaeal Cdc45/RecJ | Cdc45 | In many archaea and eukaryotes, CDC45/RecJ apparently contain inactive DHH phosphoesterase domains. The RecJ family is triplicated in euryarchaea, and some of the paralogs could be involved in repair. MCM is independently duplicated in several lineages of euryarchaea. |
Mcm | Mcm2, Mcm3, Mcm4, Mcm5, Mcm6, Mcm7 | |
Gins23 | Gins2, Gins3 | |
Gins15 | Gins1, Gins5 | |
Inactivated MCM homologa | Mcm10 | |
CMG activation factors | ||
— | RecQ/Sld2 | There is no evidence that kinases and phosphatases in archaea are directly involved in replication, although they probably regulate cell division. |
— | Treslin/Sld3 | |
— | TopBP1/Dpb11 | |
STK | CDK, DDK | |
PP2C | PP2C | |
Primases | ||
Prim1/p48 | PriS | In eukaryotes, Pol α is involved in priming by adding short DNA fragments to RNA primers. In archaea, DnaG might be involved in priming specifically on the lagging strand. |
Prim2a/p58 | PriL | |
DnaG | — | |
Polymerases | ||
PolB3 | Pol α, Pol δ, Pol ζ | No eukaryotic homologs of DP2 are known, but Zn fingers of Pol ε are apparently derived from DP2. |
PolB1 | Pol ε | |
DP1 | B subunits of Pol α, Pol δ, Pol ζ, Pol ε | |
DP2 | — | |
DNA polymerase sliding clamp and clamp loader | ||
RFCL | RFC1 | Eukaryotes have additional duplications of both RFCs and PCNA involved in checkpoint complexes (Rad27 and Rad1, Rad9, Hus1, respectively). |
RFCS | RFC2, RFC3, RFC4, RFC4 | |
PCNA | PCNA | |
Primer removal and gap closure | ||
RNase H2 | RNase II | There is a triplication of ligases (LigI, LigIII, LigIV) in eukaryotes, but only LigI is directly involved in replication. In a few Halobacteria, ATP-dependent ligase is replaced by NAD-dependent ligase. |
Fen1 | Fen1/EXO1, Rad2, Rad27 | |
Lig1 | Lig1 | |
SSB | ||
arRPA1_long | Rpa1 | In Thermoproteales, RPA is displaced by the non-homologous ThermoSSB; two short RPA forms in many euryarchaea; expansion of short RPA forms in Halobacteria. |
arRPA1_short and RPA2 | Rpa2 | |
arCOG05741a | Rpa3 |
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David M. Feliciano Angélique Bordey Luca Bonfanti 《Cold Spring Harbor perspectives in biology》2015,7(10)
Two decades after the discovery that neural stem cells (NSCs) populate some regions of the mammalian central nervous system (CNS), deep knowledge has been accumulated on their capacity to generate new neurons in the adult brain. This constitutive adult neurogenesis occurs throughout life primarily within remnants of the embryonic germinal layers known as “neurogenic sites.” Nevertheless, some processes of neurogliogenesis also occur in the CNS parenchyma commonly considered as “nonneurogenic.” This “noncanonical” cell genesis has been the object of many claims, some of which turned out to be not true. Indeed, it is often an “incomplete” process as to its final outcome, heterogeneous by several measures, including regional location, progenitor identity, and fate of the progeny. These aspects also strictly depend on the animal species, suggesting that persistent neurogenic processes have uniquely adapted to the brain anatomy of different mammals. Whereas some examples of noncanonical neurogenesis are strictly parenchymal, others also show stem cell niche-like features and a strong link with the ventricular cavities. This work will review results obtained in a research field that expanded from classic neurogenesis studies involving a variety of areas of the CNS outside of the subventricular zone (SVZ) and subgranular zone (SGZ). It will be highlighted how knowledge concerning noncanonical neurogenic areas is still incomplete owing to its regional and species-specific heterogeneity, and to objective difficulties still hampering its full identification and characterization.The central nervous system (CNS) of adult mammals is assembled during developmental neurogenesis, and its architectural specificity is maintained through a vast cohort of membrane-bound and extracellular matrix molecules (Gumbiner 1996; Bonfanti 2006). Although CNS structure is sculpted by experience-dependent synaptic plasticity at different postnatal developmental stages (critical periods) (see Sale et al. 2009) and, to a lesser extent, during adulthood (Holtmaat and Svoboda 2009), the neural networks are rather stabilized in the “mature” nervous tissue (Spolidoro et al. 2009). The differentiated cellular elements forming adult neural circuitries remain substantially unchanged in terms of their number and types, because cell renewal/addition in the CNS is very low. This situation is intuitive because connectional, neurochemical, and functional specificities are fundamental features of the mature CNS in highly complex brains, allowing specific cell types to be connected and to act in a relatively invariant way (Frotscher 1992).Since the discovery of neural stem cells (NSCs) (Reynolds and Weiss 1992), we realized that the aforementioned rules of CNS stability have a main exception in two brain regions: the forebrain subventricular zone (SVZ) (Lois and Alvarez-Buylla 1994) and the hippocampal subgranular zone (SGZ) (Gage 2000). These “adult neurogenic sites” are remnants of the embryonic germinal layers (although indirectly for the SGZ, which forms ectopically from the embryonic germinative matrix), which retain stem/progenitor cells within a special microenvironment, a “niche,” allowing and regulating NSC activity (Kriegstein and Alvarez-Buylla 2009). In addition, the areas of destination (olfactory bulb and dentate gyrus) reached by neuroblasts generated within these neurogenic sites harbor specific, not fully identified yet, environmental signals allowing the integration of young, newborn neurons. These two “canonical” sites of adult neurogenesis have been found in all animal species studied so far, including humans (reviewed in Lindsey and Tropepe 2006; Bonfanti and Ponti 2008; Kempermann 2012; Grandel and Brand 2013). Although in several classes of vertebrates including fish, amphibians, and reptiles, adult neurogenesis is widespread in many areas of the CNS (Zupanc 2006; Chapouton et al. 2007; Grandel and Brand 2013), in mammals, the vast majority of the brain and spinal cord regions out of the germinal-layer-derived neurogenic sites are commonly referred to as “nonneurogenic parenchyma” (Sohur et al. 2006; Bonfanti and Peretto 2011; Bonfanti and Nacher 2012). However, this viewpoint has changed during the last few years. New examples of cell genesis, involving both neurogenesis and gliogenesis, have been shown to occur in the so-called nonneurogenic regions of the mammalian CNS (Horner et al. 2000; Dayer et al. 2005; Kokoeva et al. 2005; Luzzati et al. 2006; Ponti et al. 2008; reviewed in Butt et al. 2005; Nishiyama et al. 2009; Migaud et al. 2010; Bonfanti and Peretto 2011), suggesting that structural plasticity involving de novo neural cell genesis could be more widespread than previously thought. Apart from their temporal persistence (some of them represent examples of delayed developmental neurogenesis, which persist postnatally; see below), neurogliogenic processes vary as to their regional localization, origin, and final outcome. In this review, “noncanonical” neurogenic processes occurring in adult mammals will be reviewed by underlining their heterogeneity across the species and their differences in intensity and outcome with respect to canonical neurogenic sites.
Open in a separate windowUnshaded rows, spontaneous (constitutive) neurogenesis; shaded rows, experimentally induced neurogenesis (growth factor infusion, lesion, etc.). No functional integration has been shown to occur in any of the studies reported here.aNeuronal differentiation of newborn cells has been well documented; in all other cases, neurogenesis has been shown only until the cell-specification step, and/or assessed with less accurate analyses (reslicing not performed, neuronal differentiation not clearly shown, very few cells shown in figures, insufficient or absent quantification).bNeurogenesis reported in this region has been denied by subsequent reports. Only a set of studies are reported; gliogenesis is not considered (data modified from Bonfanti and Peretto 2011). 相似文献
Table 1.
Main sites of noncanonical neurogenesis in the mammalian brainRats | Mice | Rabbits | Monkeys | |
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Neocortex | Gould et al. 2001 Dayer et al. 2005a Tamura et al. 2007 | Shapiro et al. 2009 | Gould et al. 1999, 2001 Bernier et al. 2002 | |
Nakatomi et al. 2002a Pencea et al. 2001 Ohira et al. 2010a | Magavi et al. 2000a Chen et al. 2004a | Vessal and Darian-Smith 2010a | ||
Corpus callosum | Pencea et al. 2001 | |||
Piriform cortexb | Pekcec et al. 2006 | Shapiro et al. 2007 | Bernier et al. 2002 | |
Olfactory tubercle | Shapiro et al. 2009 | Bedard et al. 2002b | ||
Striatum | Dayer et al. 2005a | Shapiro et al. 2009 | Luzzati et al. 2006a | Bedard et al. 2002a; 2006a |
Arvidsson et al. 2002a Pencea et al. 2001 Liu et al. 2009a | Goldowitz and Hamre 1998a Cho et al. 2007a | |||
Septum | Pencea et al. 2001 | |||
Amygdala | Shapiro et al. 2009 | Luzzati et al. 2006a | Bernier et al. 2002 | |
Hippocampus (Ammon’s horn) | Rietze et al. 2000 | |||
Nakatomi et al. 2002a | ||||
Thalamus | Pencea et al. 2001 | |||
Hypothalamus | Xu et al. 2005 | Kokoeva et al. 2007 | ||
Xu et al. 2005a Pencea et al. 2001 Matsuzaki et al. 2009 Perez-Martin et al. 2010 | Kokoeva et al. 2005a Pierce and Xu 2010 | |||
Substantia nigra | Zhao et al. 2003 Zhao and Janson Lang 2009 | |||
Zhao et al. 2003 | ||||
Cerebellum | Ponti et al. 2008a | |||
Brain stem | Bauer et al. 2005 | |||
Bauer et al. 2005 |
7.
Dynamic changes in cytosolic and nuclear Ca2+ concentration are reported to play a critical regulatory role in different aspects of skeletal muscle development and differentiation. Here we review our current knowledge of the spatial dynamics of Ca2+ signals generated during muscle development in mouse, rat, and Xenopus myocytes in culture, in the exposed myotome of dissected Xenopus embryos, and in intact normally developing zebrafish. It is becoming clear that subcellular domains, either membrane-bound or otherwise, may have their own Ca2+ signaling signatures. Thus, to understand the roles played by myogenic Ca2+ signaling, we must consider: (1) the triggers and targets within these signaling domains; (2) interdomain signaling, and (3) how these Ca2+ signals integrate with other signaling networks involved in myogenesis. Imaging techniques that are currently available to provide direct visualization of these Ca2+ signals are also described.The recognition of Ca2+ as a key regulator of muscle contraction dates back to Sydney Ringer''s seminal observations in the latter part of the 19th Century (Ringer 1883; Ringer 1886; Ringer and Buxton 1887; see reviews by Martonosi 2000; Szent-Györgyi 2004). More recently, evidence is steadily accumulating to support the proposition that Ca2+ also plays a necessary and essential role in regulating embryonic muscle development and differentiation (Flucher and Andrews 1993; Ferrari et al. 1996; Lorenzon et al. 1997; Ferrari and Spitzer 1998, 1999; Wu et al. 2000; Powell et al. 2001; Jaimovich and Carrasco 2002; Li et al. 2004; Brennan et al. 2005; Harris et al. 2005; Campbell et al. 2006; Terry et al. 2006; Fujita et al. 2007; and see reviews by Berchtold et al. 2000; Ferrari et al. 2006; Al-Shanti and Stewart 2009). What is currently lacking, however, is extensive direct visualization of the spatial dynamics of the Ca2+ signals generated by developing and differentiating muscle cells. This is especially so concerning in situ studies. The object of this article, therefore, is to review and report the current state of our understanding concerning the spatial nature of Ca2+ signaling during embryonic muscle development, especially from an in vivo perspective, and to suggest possible directions for future research. The focus of our article is embryonic skeletal muscle development because of this being an area of significant current interest. Several of the basic observations reported, however, may also be common to cardiac muscle development and in some cases to smooth muscle development. What the recent development of reliable imaging techniques has most certainly done, is to add an extra dimension of complexity to understanding the roles played by Ca2+ signaling in skeletal muscle development. For example, it is clear that membrane-bound subcellular compartments, such as the nucleus (Jaimovich and Carrasco 2002), may have endogenous Ca2+ signaling activities, as do specific cytoplasmic domains, such as the subsarcolemmal space (Campbell et al. 2006). How these Ca2+ signals interact with specific down-stream targets within their particular domain, and how they might serve to communicate information among domains, will most certainly be one of the future challenges in elucidating the Ca2+-mediated regulation of muscle development.Any methodology used to study the properties of biological molecules and how they interact during development should ideally provide spatial information, because researchers increasingly need to integrate data about the interactions that underlie a biological process (such as differentiation) with information regarding the precise location within cells or an embryo where these interactions take place. Current Ca2+ imaging techniques are beginning to provide us with this spatial information, and are thus opening up exciting new avenues of investigation in our quest to understand the signaling pathways that regulate muscle development (Animal Intact animals/Cells in culture Ca2+ reporter Reporter Loading Protocol Reference Rat 1° cultures prepared from hind limb muscle of neonatal rat pups Fluo 3-AM Cells incubated in 5.4 µM reporter for 30 min at 25°C. Jaimovich et al. 2000 Mouse Myotubes grown from C2C12 subclone of the C2 mouse muscle cell line Fluo 3-AM Incubated in 5 µM reporter plus 0.1% pluronic F-127 for 1 h at r.t. Flucher and Andrews 1993 Myotubes isolated from the intercostal muscles of E18 wild-type and RyR type 3-null mice. Fluo 3-AM Cells incubated with 4 µM for 30 min at r.t. Conklin et al. 1999b Myotubes in culture prepared from newborn mice. Fluo 3-AM Cells incubated in 10 µM for 20 min. Shirokova et al. 1999 1° cultures prepared from hind limb muscle from newborn mice. Fluo 3-AM Cells incubated in 5.4 µM reporter for 30 min at 25°C. Powell et al. 2001 Embryonic day 18 (E18) isolated diaphragm muscle fibers Fluo 4-AM Incubated in 10 µM reporter for 30 min. Chun et al. 2003 Chick Myotubes prepared from leg or breast of 11-day chick embryos Fluo 3-AM Incubated in 5 µM reporter plus 0.1% pluronic F-127 for 1 h at r.t. Flucher and Andrews 1993 Myoblasts isolated from thigh muscle of E12 embryos. Fluo 3-AM 1 mM stock was diluted 1:200 with 0.2% pluronic F-127. Cells were incubated for 60 min at r.t. in the dark. Tabata et al. 2006 Xenopus Exposed myotome in dissected embryo Fluo-3 AM Incubated dissected tissue in 10 µM reporter for 30–60 min. Ferrari and Spitzer 1999 1° myocyte cultures prepared from stage 15 Xenopus embryos. Fluo-4 AM Cells incubated in 2 µM reporter plus 0.01% pluronic F-127 for 60 min. Campbell et al. 2006 Zebrafish Intact animals Calcium green-1 dextran (10S) Reporter at 20 mM was injected into a single blastomere between the 32- and 128-cell stage. Zimprich et al. 1998 Intact animals Oregon Green 488 BAPTA dextran Single blastomeres from 32-cell stage embryos injected with reporter (i.c. 100 µM) and tetramethylrhodamine dextran (i.c. 40 µM). Ashworth et al. 2001 Intact animals Oregon Green 488 BAPTA dextran Microinjected with rhodamine dextran to give an intracellular concentration of ∼40 µM. Ashworth 2004 Intact animals Aequorin aEmbryos injected with 700 pg aeq-mRNA at the 1-cell stage and then incubated with 50 µM f-coelenterazine from the 64-cell stage. Cheung et al. 2006 Intact animals Aequorin Transgenic fish that express apoaequorin in the skeletal muscles were incubated with 50 µM f-coelenterazine from the 8-cell stage. Cheung et al. 2010