Heparan Sulfate Proteoglycans

Heparan sulfate proteoglycans are found at the cell surface and in the extracellular matrix, where they interact with a plethora of ligands. Over the last decade, new insights have emerged regarding the mechanism and biological significance of these interactions. Here, we discuss changing views on the specificity of protein–heparan sulfate binding and the activity of HSPGs as receptors and coreceptors. Although few in number, heparan sulfate proteoglycans have profound effects at the cellular, tissue, and organismal level.Heparan sulfate proteoglycans (HSPGs) are glycoproteins, with the common characteristic of containing one or more covalently attached heparan sulfate (HS) chains, a type of glycosaminoglycan (GAG) (Esko et al. 2009). Cells elaborate a relatively small set of HSPGs (∼17) that fall into three groups according to their location: membrane HSPGs, such as syndecans and glycosylphosphatidylinositol-anchored proteoglycans (glypicans), the secreted extracellular matrix HSPGs (agrin, perlecan, type XVIII collagen), and the secretory vesicle proteoglycan, serglycin (Esko et al. 1985), which allowed functional studies in the context of a cell culture model (Zhang et al. 2006). A decade later, the first HSPG mutants in a model organism (Drosophila melanogaster) were identified (Rogalski et al. 1993; Nakato et al. 1995; Häcker et al. 1997; Bellaiche et al. 1998; Lin et al. 1999), which was followed by identification of mutants in nematodes, tree frogs, zebrafish, and mice (​and3).3). HS is evolutionarily ancient and its composition has remained relatively constant from Hydra to humans (Yamada et al. 2007; Lawrence et al. 2008).

Table 1.

Heparan sulfate proteoglycans

Functional and structural differences between skinned and intact muscle preparations

Myofilaments and their associated proteins, which together constitute the sarcomeres, provide the molecular-level basis for contractile function in all muscle types. In intact muscle, sarcomere-level contraction is strongly coupled to other cellular subsystems, in particular the sarcolemmal membrane. Skinned muscle preparations (where the sarcolemma has been removed or permeabilized) are an experimental system designed to probe contractile mechanisms independently of the sarcolemma. Over the last few decades, experiments performed using permeabilized preparations have been invaluable for clarifying the understanding of contractile mechanisms in both skeletal and cardiac muscle. Today, the technique is increasingly harnessed for preclinical and/or pharmacological studies that seek to understand how interventions will impact intact muscle contraction. In this context, intrinsic functional and structural differences between skinned and intact muscle pose a major interpretational challenge. This review first surveys measurements that highlight these differences in terms of the sarcomere structure, passive and active tension generation, and calcium dependence. We then highlight the main practical challenges and caveats faced by experimentalists seeking to emulate the physiological conditions of intact muscle. Gaining an awareness of these complexities is essential for putting experiments in due perspective.

IntroductionIn striated muscle, force is generated by sarcomeres located within myocytes (Bers, 2001, 2002). The sarcomere is located within the selectively permeable cell membrane, which supports intracellular ionic homeostasis. Within this highly regulated space, sarcomere force generation is activated by dynamic changes in cytosolic Ca²⁺. The sarcomeric protein troponin C (TnC) binds to Ca²⁺, which prompts the formation of myosin cross-bridges between the sarcomere thick (myosin) and thin (actin) filaments. These myofilaments are arranged in a regular lattice oriented along the muscle fiber direction and form the main structural basis of myocyte contraction. The contraction process is regulated by many other intracellular molecules and ions, in particular Mg²⁺ and H⁺, as well as by cellular and sarcomeric morphologies.To identify the ionic and molecular mechanisms that regulate the sarcomere, it is necessary to control the chemical environment it is exposed to. The biochemistry of the sarcomere proteins can be studied using in vitro biochemistry assays. However, these fail to account for the regular structure of the sarcomere, which is important for both biochemistry and function. Alternatively, the sarcomeres can be accessed by skinning the muscle, i.e., removing the sarcolemma membrane (or making it permeable to compounds and ions), while preserving sarcomere functionality (Curtin et al., 2015). Exposing the sarcomeres to tailored ionic conditions provides a means to observe and control molecular behavior in a setting that more closely resembles native structures. After skinning, the sarcomere system is effectively isolated from the other cellular subsystems (except in some skeletal muscle experiments that remove the sarcolemma while preserving intracellular organelles and structures; Donaldson, 1985; Fill and Best, 1988; Posterino et al., 2000). This facilitates the study of contraction and its regulation separately from the sarcolemma. The central assumption of skinned muscle experiments is that the response of the sarcomeres to changes in the natural cytosol can be reproduced artificially and controllably through analogous changes in the bathing solution.In skinning protocols (typically used with skeletal muscle) where the SR is preserved, applying caffeine liberates the intracellular Ca²⁺ reserves to stimulate contraction (Donaldson, 1985). In cases where the T tubules are preserved in the skinning process, ionic substitution in the bathing solution may induce T-tubule membrane depolarization and hence Ca²⁺ release from the SR (Fill and Best, 1988). An alternative approach to releasing SR calcium is by electric-field stimulation, with the electric field applied transversely relative to the fiber direction (Posterino et al., 2000).The principal readouts of skinned-muscle experiments are contraction kinetics, adenosine triphosphatase (ATPase) activity, and generated force. Their value therefore rests on the premise that the structural integrity of the sarcomeres is preserved. Under this condition, skinned muscle may be viewed as an intermediary experimental system, straddling intact muscle and in vitro molecular experiments.Skinned preparations allow the probing of muscle behavior beyond the current reach of experiments on intact systems. In experiments where contraction is elicited by controlling the bath [Ca²⁺], the influence of “cytosolic” conditions on Ca²⁺ sensitivity, in the steady-state, is typically presented in terms of Hill-type force-[Ca²⁺] relationships, or “F-pCa,” where pCa ≡ − log₁₀[Ca²⁺]/(mol/liter). Other intracellular molecular structures that fulfill structural and mechanical roles (e.g., titin [Cazorla et al., 2001; Fukuda and Granzier, 2005; Fukuda et al., 2005; Li et al., 2016; Tonino et al., 2017] or the cytoskeleton [Roos and Brady, 1989]) can also be investigated. The controlled progression of the system from one equilibrium state to another has helped to reveal, for example, hysteresis in F-pCa, which may potentially fulfill a physiological role but would be difficult to identify in the dynamic natural system (Bers, 2001; Harrison et al., 1988). Dynamic mechanical experiments also yield insight into myofilament kinetics (Breithaupt et al., 2019; Palmer et al., 2020; Stelzer et al., 2006; Terui et al., 2010). In some (mechanical) skinning methods that preserve the T tubules, further details of the excitation–contraction coupling become experimentally accessible (Fill and Best, 1988; Posterino et al., 2000). The ability to perform protein-exchange manipulations (e.g., cardiac versus skeletal TnC; Babu et al., 1988; Gulati and Babu, 1989), to include fluorescent proteins (e.g., troponin; Brenner et al., 1999), and to perform time-resolved dynamics measurements through the flash photolysis of caged compounds (ATP [Goldman et al., 1982, 1984], inorganic phosphate [Araujo and Walker, 1996; Dantzig et al., 1992; Millar and Homsher, 1990; Tesi et al., 2000], and Ca²⁺ chelators [Luo et al., 2002; Wahr et al., 1998]) provide additional handles for probing molecular mechanisms. Overall, much of our understanding of striated muscle generally and cytosolic conditions (temperature, pH, etc.) is derived from skinned-muscle experiments (Bers, 2001).Historically, skinning has been performed in a wide array of animal species and striated muscle systems, ranging from single cells to multicellular fibers of cardiac, skeletal, and smooth muscle. Various skinning techniques have been proposed. In “mechanical” skinning, the sarcolemma is effectively peeled off (entirely or partially; Cassens et al., 1986; Endo, 1977; Trube, 1978) by microdissection (Azimi et al., 2020; Donaldson, 1985; Fabiato, 1985b; Fabiato and Fabiato, 1975, 1977, 1978a, 1978b; Fill and Best, 1988; Godt, 1974; Godt and Maughan, 1977; Jewell, 1977; Lamb and Stephenson, 2018; Matsubara and Elliott, 1972; Moisescu, 1976; Rebbeck et al., 2020), while preserving the structural integrity and function of the T tubules and the SR (Lamb and Stephenson, 1990; Posterino et al., 2000; Stephenson, 1981). However, the technique is difficult and no longer used routinely. In contrast, “chemical” skinning involves dissolving or permeabilizing the membrane by applying a chemical agent. The most common agent is Triton X-100 (Solaro et al., 1971), but alternatives include Brij (Hibberd and Jewell, 1982), lubrol (Scheld et al., 1989), glycerol, and saponin (Edes et al., 1995; Endo and Iino, 1980; Gwathmey and Hajjar, 1990; Launikonis and Stephenson, 1997; Patel et al., 2001). Chemical skinning is particularly appropriate for multicellular tissue preparations. Controlling the precise protocol and chemical agent reportedly allows the selective dissolution of the sarcolemma membrane while leaving intracellular organelles (mitochondria and SR) intact. Nonetheless, treatment with (typically 1%) Triton X-100 frees the myofibrils of contamination by mitochondrial, sarcolemmal, and SR membranes while preserving ATPase activity and sensitivity to Ca²⁺ (Solaro et al., 1971). This straightforwardness makes Triton X-100 demembranation the predominantly used technique today. Other reported skinning approaches use propionate (Reuben et al., 1971) or the Ca²⁺ chelators EGTA or EDTA (Thomas, 1960; Winegard, 1971; Miller, 1979), but the uncertainty in the underlying mechanisms has undermined the reliability of these methods (Miller, 1979). For completeness, we also mention a less used “freeze drying” approach that arguably preserves the protein content of the fibers better than chemical skinning (De Beer et al., 1992; Schiereck et al., 1993; Stienen et al., 1983).Although, for many years, skinned muscle experiments have served as an invaluable method for investigating fundamental physiology, they are increasingly inspiring more ambitious practical applications. At a practical level, live human cells are inevitably a highly scarce resource, with facilities for collecting, storing, and measuring samples often being displaced both geographically and temporally. These issues are more realistically resolved with skinned cells, which can be preserved frozen for several months (Mosqueira et al., 2019). The development of new sarcomere drugs, including omecamtiv mecarbil and mavacamten, demonstrate that the sarcomere is a viable drug target (Tsukamoto, 2019). Similarly, Ca²⁺-sensitizing drugs (which act by increasing either the sensitivity to [Ca²⁺] or the magnitude of the generated force) such as levosimendan (Edes et al., 1995), pimobendan (Fitton and Brogden, 1994; Scheld et al., 1989), sulmazole (Solaro and Rüegg, 1982), isomazole (Lues et al., 1988), and EMD-57033 (Gross et al., 1993; Lee and Allen, 1997) have all been assessed using measurements on skinned fibers. Identifying further novel sarcomere modulator compounds requires large high-throughput screening, which is unrealistic using intact muscle.There is also a growing appetite for exploiting the quantitative value of skinned muscle experiments for more direct clinical applications, such as guiding patient-specific therapies. Much of this ambition relies on the integrative power of computational models to simulate human heart mechanics based on individual patients’ data, linking sub-cellular mechanisms with systemic behavior (Niederer et al., 2019a, 2019b). Building upon basic understanding of muscle behavior, recent developments in biomedical engineering extrapolate physiological processes at the cellular and tissue levels to predict global whole-heart function. As this field continues to grow in maturity, and as model predictions allow more meaningful comparisons with clinical data, efforts are increasingly focusing on quantitatively elucidating the interdependence between cellular behavior, tissue properties, and the anatomy. The quantitative accuracy of the subsystems at all these levels therefore becomes paramount.In both of these evolving applications, the relevance and value of skinned-muscle experiments hinges on their ability to reliably emulate the intact system (Land et al., 2017; Margara et al., 2021; Mijailovich et al., 2021). Skinned-muscle experiments conducted over the past decades confirm the fidelity, in many respects, of these preparations as valid experimental models. However, they also highlight caveats and significant interpretational challenges. Gaining an awareness of these issues is becoming all the more essential to avoid misinterpretations that may have practical consequences. This review therefore aims to highlight these challenges, to help users of skinned-based measurements put them in an appropriate perspective.The present review is structured as follows. We first compare measurements of the principal physiological properties of skinned and intact muscle, highlighting similarities and discrepancies. We focus primarily on chemical skinning, and in particular Triton X-100 (the predominantly used chemical agent). We then describe practical challenges involved in conducting experiments, insofar as they impact on measurement outcomes. We conclude with a summary of recommendations and main caveats.Comparing skinned and intact muscleSkinned muscle experiments aim to reveal and controllably reproduce features of the physiological function of sarcomeres. However, notable discrepancies arise between skinned- and intact-muscle measurements of basic muscle properties that govern overall muscle function. To establish these differences rigorously at the single-cell level encounters significant methodological challenges. Although it might seem obvious that this would require doing measurements systematically on both preparation types in tandem, many early experiments were done predominantly on skinned rather than on intact cells (King et al., 2011). This stems largely from the specific challenges of noninjurious cell attachment and performing small-force measurement on intact single cells (Brady, 1991). More recently, technical developments (e.g., involving the use of flexible carbon fibers to hold the cells at opposite ends; Iribe et al., 2007; Le Guennec et al., 1990; Yasuda et al., 2001) have made these measurements more practicable. Despite these advances, however, only a fraction of studies in the literature have systematically made direct comparisons between skinned and intact systems taken from the same species under optimally similar conditions (see the selection listed in

Focus on Metabolism: Posttranslational Protein Modifications in Plant Metabolism

Posttranslational modifications (PTMs) of proteins greatly expand proteome diversity, increase functionality, and allow for rapid responses, all at relatively low costs for the cell. PTMs play key roles in plants through their impact on signaling, gene expression, protein stability and interactions, and enzyme kinetics. Following a brief discussion of the experimental and bioinformatics challenges of PTM identification, localization, and quantification (occupancy), a concise overview is provided of the major PTMs and their (potential) functional consequences in plants, with emphasis on plant metabolism. Classic examples that illustrate the regulation of plant metabolic enzymes and pathways by PTMs and their cross talk are summarized. Recent large-scale proteomics studies mapped many PTMs to a wide range of metabolic functions. Unraveling of the PTM code, i.e. a predictive understanding of the (combinatorial) consequences of PTMs, is needed to convert this growing wealth of data into an understanding of plant metabolic regulation.The primary amino acid sequence of proteins is defined by the translated mRNA, often followed by N- or C-terminal cleavages for preprocessing, maturation, and/or activation. Proteins can undergo further reversible or irreversible posttranslational modifications (PTMs) of specific amino acid residues. Proteins are directly responsible for the production of plant metabolites because they act as enzymes or as regulators of enzymes. Ultimately, most proteins in a plant cell can affect plant metabolism (e.g. through effects on plant gene expression, cell fate and development, structural support, transport, etc.). Many metabolic enzymes and their regulators undergo a variety of PTMs, possibly resulting in changes in oligomeric state, stabilization/degradation, and (de)activation (Huber and Hardin, 2004), and PTMs can facilitate the optimization of metabolic flux. However, the direct in vivo consequence of a PTM on a metabolic enzyme or pathway is frequently not very clear, in part because it requires measurements of input and output of the reactions, including flux through the enzyme or pathway. This Update will start out with a short overview on the major PTMs observed for each amino acid residue (PTMs, including determination of the localization within proteins (i.e. the specific residues) and occupancy. Challenges in dealing with multiple PTMs per protein and cross talk between PTMs will be briefly outlined. We then describe the major physiological PTMs observed in plants as well as PTMs that are nonenzymatically induced during sample preparation (PTMs, in particular for enzymes in primary metabolism (Calvin cycle, glycolysis, and respiration) and the C4 shuttle accommodating photosynthesis in C4 plants (PTMs observed in plants

Variation in Adult Plant Phenotypes and Partitioning among Seed and Stem-Borne Roots across Brachypodium distachyon Accessions to Exploit in Breeding Cereals for Well-Watered and Drought Environments

Seedling roots enable plant establishment. Their small phenotypes are measured routinely. Adult root systems are relevant to yield and efficiency, but phenotyping is challenging. Root length exceeds the volume of most pots. Field studies measure partial adult root systems through coring or use seedling roots as adult surrogates. Here, we phenotyped 79 diverse lines of the small grass model Brachypodium distachyon to adults in 50-cm-long tubes of soil with irrigation; a subset of 16 lines was droughted. Variation was large (total biomass, ×8; total root length [TRL], ×10; and root mass ratio, ×6), repeatable, and attributable to genetic factors (heritabilities ranged from approximately 50% for root growth to 82% for partitioning phenotypes). Lines were dissected into seed-borne tissues (stem and primary seminal axile roots) and stem-borne tissues (tillers and coleoptile and leaf node axile roots) plus branch roots. All lines developed one seminal root that varied, with branch roots, from 31% to 90% of TRL in the well-watered condition. With drought, 100% of TRL was seminal, regardless of line because nodal roots were almost always inhibited in drying topsoil. Irrigation stimulated nodal roots depending on genotype. Shoot size and tillers correlated positively with roots with irrigation, but partitioning depended on genotype and was plastic with drought. Adult root systems of B. distachyon have genetic variation to exploit to increase cereal yields through genes associated with partitioning among roots and their responsiveness to irrigation. Whole-plant phenotypes could enhance gain for droughted environments because root and shoot traits are coselected.Adult plant root systems are relevant to the size and efficiency of seed yield. They supply water and nutrients for the plant to acquire biomass, which is positively correlated to the harvest index (allocation to seed grain), and the stages of flowering and grain development. Modeling in wheat (Triticum aestivum) suggested that an extra 10 mm of water absorbed by such adult root systems during grain filling resulted in an increase of approximately 500 kg grain ha⁻¹ (Manschadi et al., 2006). This was 25% above the average annual yield of wheat in rain-fed environments of Australia. This number was remarkably close to experimental data obtained in the field in Australia (Kirkegaard et al., 2007). Together, these modeling and field experiments have shown that adult root systems are critical for water absorption and grain yield in cereals, such as wheat, emphasizing the importance of characterizing adult root systems to identify phenotypes for productivity improvements.Most root phenotypes, however, have been described for seedling roots. Seedling roots are essential for plant establishment, and hence, the plant’s potential to set seed. For technical reasons, seedlings are more often screened than adult plants because of the ease of handling smaller plants and the high throughput. Seedling-stage phenotyping may also improve overall reproducibility of results because often, growth media are soil free. Seedling soil-free root phenotyping conditions are well suited to dissecting fine and sensitive mechanisms, such as lateral root initiation (Casimiro et al., 2003; Péret et al., 2009a, 2009b). A number of genes underlying root processes have been identified or characterized using seedlings, notably with the dicotyledonous models Arabidopsis (Arabidopsis thaliana; Mouchel et al., 2004; Fitz Gerald et al., 2006; Yokawa et al., 2013) and Medicago truncatula (Laffont et al., 2010) and the cereals maize (Zea mays; Hochholdinger et al., 2001) and rice (Oryza sativa; Inukai et al., 2005; Kitomi et al., 2008).Extrapolation from seedling to adult root systems presents major questions (Hochholdinger and Zimmermann, 2008; Chochois et al., 2012; Rich and Watt, 2013). Are phenotypes in seedling roots present in adult roots given developmental events associated with aging? Is expression of phenotypes correlated in seedling and adult roots if time compounds effects of growth rates and growth conditions on roots? Watt et al. (2013) showed in wheat seedlings that root traits in the laboratory and field correlated positively but that neither correlated with adult root traits in the field. Factors between seedling and adult roots seemed to be differences in developmental stage and the time that growing roots experience the environment.Seedling and adult root differences may be larger in grasses than dicotyledons. Grass root systems have two developmental components: seed-borne (seminal) roots, of which a number emerge at germination and continue to grow and branch throughout the plant life, and stem-borne (nodal or adventitious) roots, which emerge from around the three-leaf stage and continue to emerge, grow, and branch throughout the plant life. Phenotypes and traits of adult root systems of grasses, which include the major cereal crops wheat, rice, and maize, are difficult to predict in seedling screens and ideally identified from adult root systems first (Gamuyao et al., 2012).Phenotyping of adult roots is possible in the field using trenches (Maeght et al., 2013) or coring (Wasson et al., 2014). A portion of the root system is captured with these methods. Alternatively, entire adult root systems can be contained within pots dug into the ground before sowing. These need to be large; field wheat roots, for example, can reach depths greater than 1.5 m depending on genotype and environment. This method prevents root-root interactions that occur under normal field sowing of a plant canopy and is also a compromise.A solution to the problem of phenotyping adult cereal root systems is a model for monocotyledon grasses: Brachypodium distachyon. B. distachyon is a small-stature grass with a small genome that is fully sequenced (Vogel et al., 2010). It has molecular tools equivalent to those available in Arabidopsis (Draper et al., 2001; Brkljacic et al., 2011; Mur et al., 2011). The root system of B. distachyon reference line Bd21 is more similar to wheat than other model and crop grasses (Watt et al., 2009). It has a seed-borne primary seminal root (PSR) that emerges from the embryo at seed germination and multiple stem-borne coleoptile node axile roots (CNRs) and leaf node axile roots (LNRs), also known as crown roots or adventitious roots, that emerge at about three leaves through to grain development. Branch roots emerge from all root types. There are no known anatomical differences between root types of wheat and B. distachyon (Watt et al., 2009). In a recent study, we report postflowering root growth in B. distachyon line Bd21-3, showing that this model can be used to answer questions relevant to the adult root systems of grasses (Chochois et al., 2012).In this study, we used B. distachyon to identify adult plant phenotypes related to the partitioning among seed-borne and stem-borne shoots and roots for the genetic improvement of well-watered and droughted cereals (Fig. 1; Krassovsky, 1926; Navara et al., 1994), nitrogen, phosphorus (Tennant, 1976; Brady et al., 1995), oxygen (Wiengweera and Greenway, 2004), soil hardness (Acuna et al., 2007), and microorganisms (Sivasithamparam et al., 1978). Of note is the study by Krassovsky (1926), which was the first, to our knowledge, to show differences in function related to water. Krassovsky (1926) showed that seminal roots of wheat absorbed almost 2 times the water as nodal roots per unit dry weight but that nodal roots absorbed a more diluted nutrient solution than seminal roots. Krassovsky (1926) also showed by removing seminal or nodal roots as they emerged that “seminal roots serve the main stem, while nodal roots serve the tillers” (Krassovsky, 1926). Volkmar (1997) showed, more recently, in wheat that nodal and seminal roots may sense and respond to drought differently. In millet (Pennisetum glaucum) and sorghum (Sorghum bicolor), Rostamza et al. (2013) found that millet was able to grow nodal roots in a dryer soil than sorghum, possibly because of shoot and root vigor.Open in a separate windowFigure 1.B. distachyon plant scanned at the fourth leaf stage, with the root and shoot phenotypes studied indicated. Supplemental Table S1.

Arabidopsis LON2 Is Necessary for Peroxisomal Function and Sustained Matrix Protein Import

Relatively little is known about the small subset of peroxisomal proteins with predicted protease activity. Here, we report that the peroxisomal LON2 (At5g47040) protease facilitates matrix protein import into Arabidopsis (Arabidopsis thaliana) peroxisomes. We identified T-DNA insertion alleles disrupted in five of the nine confirmed or predicted peroxisomal proteases and found only two—lon2 and deg15, a mutant defective in the previously described PTS2-processing protease (DEG15/At1g28320)—with phenotypes suggestive of peroxisome metabolism defects. Both lon2 and deg15 mutants were mildly resistant to the inhibitory effects of indole-3-butyric acid (IBA) on root elongation, but only lon2 mutants were resistant to the stimulatory effects of IBA on lateral root production or displayed Suc dependence during seedling growth. lon2 mutants displayed defects in removing the type 2 peroxisome targeting signal (PTS2) from peroxisomal malate dehydrogenase and reduced accumulation of 3-ketoacyl-CoA thiolase, another PTS2-containing protein; both defects were not apparent upon germination but appeared in 5- to 8-d-old seedlings. In lon2 cotyledon cells, matrix proteins were localized to peroxisomes in 4-d-old seedlings but mislocalized to the cytosol in 8-d-old seedlings. Moreover, a PTS2-GFP reporter sorted to peroxisomes in lon2 root tip cells but was largely cytosolic in more mature root cells. Our results indicate that LON2 is needed for sustained matrix protein import into peroxisomes. The delayed onset of matrix protein sorting defects may account for the relatively weak Suc dependence following germination, moderate IBA-resistant primary root elongation, and severe defects in IBA-induced lateral root formation observed in lon2 mutants.Peroxisomes are single-membrane-bound organelles found in most eukaryotes. Peroxin (PEX) proteins are necessary for various aspects of peroxisome biogenesis, including matrix protein import (for review, see Distel et al., 1996; Schrader and Fahimi, 2008). Most matrix proteins are imported into peroxisomes from the cytosol using one of two targeting signals, a C-terminal type 1 peroxisome-targeting signal (PTS1) or a cleavable N-terminal type 2 peroxisome-targeting signal (PTS2) (Reumann, 2004). PTS1- and PTS2-containing proteins are bound in the cytosol by soluble matrix protein receptors, escorted to the peroxisome membrane docking complex, and translocated into the peroxisome matrix (for review, see Platta and Erdmann, 2007). Once in the peroxisome, many matrix proteins participate in metabolic pathways, such as β-oxidation, hydrogen peroxide decomposition, and photorespiration (for review, see Gabaldon et al., 2006; Poirier et al., 2006).In addition to metabolic enzymes, several proteases are found in the peroxisome matrix. Only one protease, DEG15/Tysnd1, has a well-defined role in peroxisome biology. The rat Tysnd1 protease removes the targeting signal after PTS2-containing proteins enter the peroxisome and also processes certain PTS1-containing β-oxidation enzymes (Kurochkin et al., 2007). Similarly, the Arabidopsis (Arabidopsis thaliana) Tysnd1 homolog DEG15 (At1g28320) is a peroxisomal Ser protease that removes PTS2 targeting signals (Helm et al., 2007; Schuhmann et al., 2008).In contrast with DEG15, little is known about the other eight Arabidopsis proteins that are annotated as proteases in the AraPerox database of putative peroxisomal proteins (Reumann et al., 2004; Carter et al., 2004; Shimaoka et al., 2004), which, in combination with the minor PTS found in both of these predicted proteases (Reumann, 2004), suggests that these enzymes may not be peroxisomal. Along with DEG15, only two of the predicted peroxisomal proteases, an M16 metalloprotease (At2g41790), which we have named PXM16 for peroxisomal M16 protease, and a Lon-related protease (At5g47040/LON2; Ostersetzer et al., 2007), are found in the proteome of peroxisomes purified from Arabidopsis suspension cells (Eubel et al., 2008). DEG15 and LON2 also have been validated as peroxisomally targeted using GFP fusions (Ostersetzer et al., 2007; Schuhmann et al., 2008).

Table I.

Putative Arabidopsis proteases predicted or demonstrated to be peroxisomal

Visualization of Ca2+ Signaling During Embryonic Skeletal Muscle Formation in Vertebrates

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 (AnimalIntact animals/Cells in cultureCa²⁺ reporterReporter Loading ProtocolReferenceRat1° cultures prepared from hind limb muscle of neonatal rat pupsFluo 3-AMCells incubated in 5.4 µM reporter for 30 min at 25°C.Jaimovich et al. 2000MouseMyotubes grown from C2C12 subclone of the C2 mouse muscle cell lineFluo 3-AMIncubated in 5 µM reporter plus 0.1% pluronic F-127 for 1 h at r.t.Flucher and Andrews 1993Myotubes isolated from the intercostal muscles of E18 wild-type and RyR type 3-null mice.Fluo 3-AMCells incubated with 4 µM for 30 min at r.t.Conklin et al. 1999bMyotubes in culture prepared from newborn mice.Fluo 3-AMCells incubated in 10 µM for 20 min.Shirokova et al. 19991° cultures prepared from hind limb muscle from newborn mice.Fluo 3-AMCells incubated in 5.4 µM reporter for 30 min at 25°C.Powell et al. 2001Embryonic day 18 (E18) isolated diaphragm muscle fibersFluo 4-AMIncubated in 10 µM reporter for 30 min.Chun et al. 2003ChickMyotubes prepared from leg or breast of 11-day chick embryosFluo 3-AMIncubated in 5 µM reporter plus 0.1% pluronic F-127 for 1 h at r.t.Flucher and Andrews 1993Myoblasts isolated from thigh muscle of E12 embryos.Fluo 3-AM1 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. 2006XenopusExposed myotome in dissected embryoFluo-3 AMIncubated dissected tissue in 10 µM reporter for 30–60 min.Ferrari and Spitzer 19991° myocyte cultures prepared from stage 15 Xenopus embryos.Fluo-4 AMCells incubated in 2 µM reporter plus 0.01% pluronic F-127 for 60 min.Campbell et al. 2006ZebrafishIntact animalsCalcium green-1 dextran (10S)Reporter at 20 mM was injected into a single blastomere between the 32- and 128-cell stage.Zimprich et al. 1998Intact animalsOregon Green 488 BAPTA dextranSingle blastomeres from 32-cell stage embryos injected with reporter (i.c. 100 µM) and tetramethylrhodamine dextran (i.c. 40 µM).Ashworth et al. 2001Intact animalsOregon Green 488 BAPTA dextranMicroinjected with rhodamine dextran to give an intracellular concentration of ∼40 µM.Ashworth 2004Intact animalsAequorin^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. 2006Intact animalsAequorinTransgenic fish that express apoaequorin in the skeletal muscles were incubated with 50 µM f-coelenterazine from the 8-cell stage.Cheung et al. 2010Open in a separate window^aExpression of aequorin was ubiquitous but it was suggested that the Ca²⁺ signals visualized in the trunk at the approximately 8–20-somite stage and at ∼47 hpf might play a role in muscle development.     

Evolution and Function of the Plant Cell Wall Synthesis-Related Glycosyltransferase Family 8

Carbohydrate-active enzyme glycosyltransferase family 8 (GT8) includes the plant galacturonosyltransferase1-related gene family of proven and putative α-galacturonosyltransferase (GAUT) and GAUT-like (GATL) genes. We computationally identified and investigated this family in 15 fully sequenced plant and green algal genomes and in the National Center for Biotechnology Information nonredundant protein database to determine the phylogenetic relatedness of the GAUTs and GATLs to other GT8 family members. The GT8 proteins fall into three well-delineated major classes. In addition to GAUTs and GATLs, known or predicted to be involved in plant cell wall biosynthesis, class I also includes a lower plant-specific GAUT and GATL-related (GATR) subfamily, two metazoan subfamilies, and proteins from other eukaryotes and cyanobacteria. Class II includes galactinol synthases and plant glycogenin-like starch initiation proteins that are not known to be directly involved in cell wall synthesis, as well as proteins from fungi, metazoans, viruses, and bacteria. Class III consists almost entirely of bacterial proteins that are lipooligo/polysaccharide α-galactosyltransferases and α-glucosyltransferases. Sequence motifs conserved across all GT8 subfamilies and those specific to plant cell wall-related GT8 subfamilies were identified and mapped onto a predicted GAUT1 protein structure. The tertiary structure prediction identified sequence motifs likely to represent key amino acids involved in catalysis, substrate binding, protein-protein interactions, and structural elements required for GAUT1 function. The results show that the GAUTs, GATLs, and GATRs have a different evolutionary origin than other plant GT8 genes, were likely acquired from an ancient cyanobacterium (Synechococcus) progenitor, and separate into unique subclades that may indicate functional specialization.Plant cell walls are composed of three principal types of polysaccharides: cellulose, hemicellulose, and pectin. Studying the biosynthesis and degradation of these biopolymers is important because cell walls have multiple roles in plants, including providing structural support to cells and defense against pathogens, serving as cell-specific developmental and differentiation markers, and mediating or facilitating cell-cell communication. In addition to their important roles within plants, cell walls also have many economic uses in human and animal nutrition and as sources of natural textile fibers, paper and wood products, and components of fine chemicals and medicinal products. The study of the biosynthesis and biodegradation of plant cell walls has become even more significant because cell walls are the major components of biomass (Mohnen et al., 2008), which is the most promising renewable source for the production of biofuels and biomaterials (Ragauskas et al., 2006; Pauly and Keegstra, 2008). Analyses of fully sequenced plant genomes have revealed that they encode hundreds or even thousands of carbohydrate-active enzymes (CAZy; Henrissat et al., 2001; Yokoyama and Nishitani, 2004; Geisler-Lee et al., 2006). Most of these CAZy enzymes (Cantarel et al., 2009) are glycosyltransferases (GTs) or glycoside hydrolases, which are key players in plant cell wall biosynthesis and modification (Cosgrove, 2005).The CAZy database is classified into 290 protein families (www.cazy.org; release of September 2008), of which 92 are GT families (Cantarel et al., 2009). A number of the GT families have been previously characterized to be involved in plant cell wall biosynthesis. For example, the GT2 family is known to include cellulose synthases and some hemicellulose backbone synthases (Lerouxel et al., 2006), such as mannan synthases (Dhugga et al., 2004; Liepman et al., 2005), putative xyloglucan synthases (Cocuron et al., 2007), and mixed linkage glucan synthases (Burton et al., 2006). With respect to the synthesis of xylan, a type of hemicellulose, four Arabidopsis (Arabidopsis thaliana) proteins from the GT43 family, irregular xylem 9 (IRX9), IRX14, IRX9-L, and IRX14-L, and two proteins from the GT47 family, IRX10 and IRX10-L, are candidates (York and O''Neill, 2008) for glucuronoxylan backbone synthases (Brown et al., 2007, 2009; Lee et al., 2007a; Peña et al., 2007; Wu et al., 2009). In addition, three proteins have been implicated in the synthesis of an oligosaccharide thought to act either as a primer or terminator in xylan synthesis (Peña et al., 2007): two from the GT8 family (IRX8/GAUT12 [Persson et al., 2007] and PARVUS/GATL1 [Brown et al., 2007; Lee et al., 2007b]) and one from the GT47 family (FRA8/IRX7 [Zhong et al., 2005]).The GT families involved in the biosynthesis of pectins have been relatively less studied until recently. In 2006, a gene in CAZy family GT8 was shown to encode a functional homogalacturonan α-galacturonosyltransferase, GAUT1 (Sterling et al., 2006). GAUT1 belongs to a 25-member gene family in Arabidopsis, the GAUT1-related gene family, that includes two distinct but closely related families, the galacturonosyltransferase (GAUT) genes and the galacturonosyltransferase-like (GATL) genes (Sterling et al., 2006). Another GAUT gene, GAUT8/QUA1, has been suggested to be involved in pectin and/or xylan synthesis, based on the phenotypes of plant lines carrying mutations in this gene (Bouton et al., 2002; Orfila et al., 2005). It has further been suggested that multiple members of the GT8 family are galacturonosyltransferases involved in pectin and/or xylan biosynthesis (Mohnen, 2008; Caffall and Mohnen, 2009; Caffall et al., 2009).Aside from the 25 GAUT and GATL genes, Arabidopsis has 16 other family GT8 genes, according to the CAZy database, which do not seem to have the conserved sequence motifs found in GAUTs and GATLs: HxxGxxKPW and GLG (Sterling et al., 2006). Eight of these 16 genes are annotated as galactinol synthase (GolS) by The Arabidopsis Information Resource (TAIR; www.arabidopsis.org), and three of these AtGolS enzymes have been implicated in the synthesis of raffinose family oligosaccharides that are associated with stress tolerance (Taji et al., 2002). The other eight Arabidopsis GT8 genes are annotated as plant glycogenin-like starch initiation proteins (PGSIPs) in TAIR. PGSIPs have been proposed to be involved in the synthesis of primers necessary for starch biosynthesis (Chatterjee et al., 2005). Hence, the GT8 family is a protein family consisting of enzymes with very distinct proven and proposed functions. Indeed, a suggestion has been made to split the GT8 family into two groups (Sterling et al., 2006), namely, the cell wall biosynthesis-related genes (GAUTs and GATLs) and the non-cell wall synthesis-related genes (GolSs and PGSIPs).We are interested in further defining the functions of the GAUT and GATL proteins in plants, in particular their role(s) in plant cell wall synthesis. The apparent disparate functions of the GT8 family (i.e. the GAUTs and GATLs as proven and putative plant cell wall polysaccharide biosynthetic α-galacturonosyltransferases, the eukaryotic GolSs as α-galactosyltransferases that synthesize the first step in the synthesis of the oligosaccharides stachyose and raffinose, the putative PGSIPs, and the large bacterial GT8 family of diverse α-glucosyltransferases and α-galactosyltransferases involved in lipopolysaccharide and lipooligosaccharide synthesis) indicate that the GT8 family members are involved in several unique types of glycoconjugate and glycan biosynthetic processes (Yin et al., 2010). This observation led us to ask whether any of the GT8 family members are sufficiently closely related to GAUT and GATL genes to be informative regarding GAUT or GATL biosynthetic function(s) and/or mechanism(s).To investigate the relatedness of the members of the GT8 gene family, we carried out a detailed phylogenetic analysis of the entire GT8 family in 15 completely sequenced plant and green algal genomes (AbbreviationCladeSpeciesGenome PublishedDownloaded frommpcGreen algaeMicromonas pusilla CCMP1545Worden et al. (2009)JGI version 2.0mprGreen algaeMicromonas strain RCC299Worden et al. (2009)JGI version 2.0olGreen algaeOstreococcus lucimarinusPalenik et al. (2007)JGI version 1.0otGreen algaeOstreococcus tauriDerelle et al. (2006)JGI version 1.0crGreen algaeChlamydomonas reinhardtiiMerchant et al. (2007)JGI version 3.0vcGreen algaeVolvox carteri f. nagariensisNoJGI version 1.0ppMossPhyscomitrella patens ssp. patensRensing et al. (2008)JGI version 1.1smSpike mossSelaginella moellendorffiiNoJGI version 1.0ptDicotPopulus trichocarpaTuskan et al. (2006)JGI version 1.1atDicotArabidopsis thalianaArabidopsis Genome Initiative (2000)TAIR version 9.0vvDicotVitis viniferaJaillon et al. (2007)http://www.genoscope.cns.fr/gmDicotGlycine maxSchmutz et al. (2010)JGI version 1.0osMonocotOryza sativaGoff et al. (2002); Yu et al. (2002)TIGR version 6.1sbMonocotSorghum bicolorPaterson et al. (2009)JGI version 1.0bdMonocotBrachypodium distachyonVogel et al. (2010)JGI version 1.0Open in a separate window     

Retrograde Intraflagellar Transport Mutants Identify Complex A Proteins With Multiple Genetic Interactions in Chlamydomonas reinhardtii

The intraflagellar transport machinery is required for the assembly of cilia. It has been investigated by biochemical, genetic, and computational methods that have identified at least 21 proteins that assemble into two subcomplexes. It has been hypothesized that complex A is required for retrograde transport. Temperature-sensitive mutations in FLA15 and FLA17 show defects in retrograde intraflagellar transport (IFT) in Chlamydomonas. We show that IFT144 and IFT139, two complex A proteins, are encoded by FLA15 and FLA17, respectively. The fla15 allele is a missense mutation in a conserved cysteine and the fla17 allele is an in-frame deletion of three exons. The flagellar assembly defect of each mutant is rescued by the respective transgenes. In fla15 and fla17 mutants, bulges form in the distal one-third of the flagella at the permissive temperature and this phenotype is also rescued by the transgenes. These bulges contain the complex B component IFT74/72, but not α-tubulin or p28, a component of an inner dynein arm, which suggests specificity with respect to the proteins that accumulate in these bulges. IFT144 and IFT139 are likely to interact with each other and other proteins on the basis of three distinct genetic tests: (1) Double mutants display synthetic flagellar assembly defects at the permissive temperature, (2) heterozygous diploid strains exhibit second-site noncomplemention, and (3) transgenes confer two-copy suppression. Since these tests show different levels of phenotypic sensitivity, we propose they illustrate different gradations of gene interaction between complex A proteins themselves and with a complex B protein (IFT172).CILIA and flagella are microtubule-based organelles that are found on most mammalian cells. They provide motility to cells and participate in many sensory processes. Defects in or loss of cilia/flagella cause a variety of human diseases that include polycystic kidney disease, retinal degeneration, infertility, obesity, respiratory defects, left–right axis determination, and polydactyly (Fliegauf et al. 2007). Mouse mutants demonstrate that cilia are essential for viability, neural tube closure, and bone development (Eggenschwiler and Anderson 2007; Fliegauf et al. 2007). Cilia and flagella are also present in protists, algae, moss, and some fungi.The assembly and maintenance of cilia and flagella require intraflagellar transport (IFT) (Kozminski et al. 1995). IFT involves the movement of 100- to 200-nm-long protein particles from the basal body located in the cell body to the tip of the flagella using the heterotrimeric kinesin-2 (anterograde movement) (Kozminski et al. 1995) and movement back to the cell body (retrograde movement) using the cytoplasmic dynein complex (Pazour et al. 1999; Porter et al. 1999). IFT particles change their direction of movement as well as their size, speed, and frequency at the ends of the flagella as they switch from anterograde to retrograde movement (Iomini et al. 2001). Biochemical isolation of IFT particles reveals that they are composed of at least 16 proteins and that these particles can be dissociated into two complexes in vitro by changing the salt concentration (Cole et al. 1998; Piperno et al. 1998). Recent genetic and bioinformatics analysis adds at least 7 more proteins to the IFT particle (Follit et al. 2009) (Eggenschwiler and Anderson 2007).

TABLE 1

Proteins and gene names for the intraflagellar transport particles in Chlamydomonas, C. elegans, and mouse

Noncanonical Sites of Adult Neurogenesis in the Mammalian Brain

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.

Table 1.

Main sites of noncanonical neurogenesis in the mammalian brain