Uniform nomenclature for the mitochondrial contact site and cristae organizing system

The mitochondrial inner membrane contains a large protein complex that functions in inner membrane organization and formation of membrane contact sites. The complex was variably named the mitochondrial contact site complex, mitochondrial inner membrane organizing system, mitochondrial organizing structure, or Mitofilin/Fcj1 complex. To facilitate future studies, we propose to unify the nomenclature and term the complex “mitochondrial contact site and cristae organizing system” and its subunits Mic10 to Mic60.Mitochondria possess two membranes of different architecture and function (Palade, 1952; Hackenbrock, 1968). Both membranes work together for essential shared functions, such as protein import (Schatz, 1996; Neupert and Herrmann, 2007; Chacinska et al., 2009). The outer membrane harbors machinery that controls the shape of the organelle and is crucial for the communication of mitochondria with the rest of the cell. The inner membrane harbors the complexes of the respiratory chain, the F₁F_o-ATP synthase, numerous metabolite carriers, and enzymes of mitochondrial metabolism. It consists of two domains: the inner boundary membrane, which is adjacent to the outer membrane, and invaginations of different shape, termed cristae (Werner and Neupert, 1972; Frey and Mannella, 2000; Hoppins et al., 2007; Pellegrini and Scorrano, 2007; Zick et al., 2009; Davies et al., 2011). Tubular openings, termed crista junctions (Perkins et al., 1997), connect inner boundary membrane and cristae membranes (Fig. 1, A and B). Respiratory chain complexes and the F₁F_o-ATP synthase are preferentially located in the cristae membranes, whereas preprotein translocases are enriched in the inner boundary membrane (Vogel et al., 2006; Wurm and Jakobs, 2006; Davies et al., 2011). Contact sites between outer membrane and inner boundary membrane promote import of preproteins, metabolite channeling, lipid transport, and membrane dynamics (Frey and Mannella, 2000; Sesaki and Jensen, 2004; Hoppins et al., 2007, 2011; Neupert and Herrmann, 2007; Chacinska et al., 2009; Connerth et al., 2012; van der Laan et al., 2012).Open in a separate windowFigure 1.MICOS complex. (A) The MICOS complex (hypothetical model), previously also termed MINOS, MitOS, or Mitofilin/Fcj1 complex, is required for maintenance of the characteristic architecture of the mitochondrial inner membrane (IM) and forms contact sites with the outer membrane (OM). In budding yeast, six subunits of MICOS have been identified. All subunits are exposed to the intermembrane space (IMS), five are integral inner membrane proteins (Mic10, Mic12, Mic26, Mic27, and Mic60), and one is a peripheral inner membrane protein (Mic19). Mic26 is related to Mic27; however, mic26Δ yeast cells show considerably less severe defects of mitochondrial inner membrane architecture than mic27Δ cells (Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011). The MICOS complex of metazoa additionally contains Mic25, which is related to Mic19, yet subunits corresponding to Mic12 and Mic26 have not been identified so far. MICOS subunits that have been conserved in most organisms analyzed are indicated by bold boundary lines. (B, top) Wild-type architecture of the mitochondrial inner membrane with crista junctions and cristae. (bottom) This architecture is considerably altered in micos mutant mitochondria: most cristae membranes are detached from the inner boundary membrane and form internal membrane stacks. In some micos mutants (deficiency of mammalian Mic19 or Mic25), a loss of cristae membranes was observed (Darshi et al., 2011; An et al., 2012). Figure by M. Bohnert (Institute of Biochemistry and Molecular Biology, University of Freiburg, Freiburg, Germany).To understand the complex architecture of mitochondria, it will be crucial to identify the molecular machineries that control the interaction between mitochondrial outer and inner membranes and the characteristic organization of the inner membrane. A convergence of independent studies led to the identification of a large heterooligomeric protein complex of the mitochondrial inner membrane conserved from yeast to humans that plays crucial roles in the maintenance of crista junctions, inner membrane architecture, and formation of contact sites to the outer membrane (Fig. 1 A). Several names were used by different research groups to describe the complex, including mitochondrial contact site (MICOS) complex, mitochondrial inner membrane organizing system (MINOS), mitochondrial organizing structure (MitOS), Mitofilin complex, or Fcj1 (formation of crista junction protein 1) complex (Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012). Mitofilin, also termed Fcj1, was the first component identified (Icho et al., 1994; Odgren et al., 1996; Gieffers et al., 1997; John et al., 2005) and was observed enriched at crista junctions (Rabl et al., 2009). Mutants of Mitofilin/Fcj1 as well as of other MICOS/MINOS/MitOS subunits show a strikingly altered inner membrane architecture. They lose crista junctions and contain large internal membrane stacks, the respiratory activity is reduced, and mitochondrial DNA nucleoids are altered (Fig. 1 B; John et al., 2005; Hess et al., 2009; Rabl et al., 2009; Mun et al., 2010; Harner et al., 2011; Head et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012; Itoh et al., 2013). It has been reported that the complex interacts with a variety of outer membrane proteins, such as channel proteins and components of the protein translocases and mitochondrial fusion machines, and defects impair the biogenesis of mitochondrial proteins (Xie et al., 2007; Darshi et al., 2011; Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012; An et al., 2012; Bohnert et al., 2012; Körner et al., 2012; Ott et al., 2012; Zerbes et al., 2012; Jans et al., 2013; Weber et al., 2013). The MICOS/MINOS/MitOS/Mitofilin/Fcj1 complex thus plays crucial roles in mitochondrial architecture, dynamics, and biogenesis. However, communication of results in this rapidly developing field has been complicated by several different nomenclatures used for the complex as well as for its subunits (

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.

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 ReferenceSystemIntactSkinning method[Mg²⁺] (mM)Ionic strength (mM)pH Reuben et al. (1971) CrayfishEGTA-3007.0 Winegard (1971) Frog cardiacEDTA1-6.5–7.0 Matsubara and Elliott (1972) Frog skeletalXDissection1-7.0 Godt (1974) Frog skeletalDissection51507.3 Wood et al. (1975) Human skeletalEGTA2–4-7.0 Moisescu (1976) Frog skeletalDissection11507.1 Godt and Maughan (1977) Frog skeletalXDissection31507.0 Best et al. (1977) Rat cardiacHomogenization0.05, 11507.0 Trube (1978) Mouse cardiacDissection (partial)41327.0 Gordon (1978) Rabbit smoothTriton X-1001.0–6.91307.0 Stienen et al. (1983) Frog skeletalFreeze drying1.11607.0Fabiato and Fabiato (1975, 1978a, 1978b)Rat cardiacDissection0.321607.0 Fabiato and Fabiato (1978a) Frog skeletalDissection0.321607.0 Fabiato (1981) Rat cardiacXEGTA11607.1 Fabiato (1981) Rabbit cardiacXEGTA11607.1 Fabiato (1985b) Canine cardiacDissection31707.1 Hibberd and Jewell (1982) Rat cardiacBrij-580.32007.0Solaro et al. (1971, 1976); Solaro and Rüegg (1982)Canine cardiacTriton X-100Var1007.0 Donaldson (1985) Rabbit skeletalDissection11507.0 Kentish et al. (1986) Rat cardiacXTriton X-10032007.0 Fill and Best (1988) Frog skeletalDissection11507.0 Lues et al. (1988) Various cardiacTriton X-100-1406.7 Roos and Brady (1989) Rat cardiacXTriton X-100-1607.1 Scheld et al. (1989) Human cardiacLubrol PX-1406.7 Harrison and Bers (1989) Rabbit cardiacTriton X-1002.2-7.0 Lamb and Stephenson (1990) Toad skeletalDissection1-7.10 Gwathmey and Hajjar (1990) Human cardiacXSaponin31607.1 Sweitzer and Moss (1990) Rat cardia, rabbit skeletalTriton X-10011807.0 Millar and Homsher (1990) Rabbit skeletalEGTA12007.1 De Beer et al. (1992) Rabbit skeletalFreeze drying--- Gross et al. (1993) Guinea pig cardiacTriton X-100--7.4 Gao et al. (1994) Rat cardiacXTriton X-1001.2-7.0 Wolff et al. (1995a) Canine cardiacTriton X-10011807.0 Edes et al. (1995) Guinea pig cardiacSaponin-1607.4 Araujo and Walker (1996) Rat cardiacTriton X-1001180- Allen et al. (2000) Rat cardiacTriton X-1001–81507.0 Posterino et al. (2000) Rat skeletalDissection1-7.1 Irving et al. (2000) Rat trabeculaeXTriton X-100-2007.35 Patel et al. (2001) Mouse cardiacSaponin + Triton X-100-1807.0 Konhilas et al. (2002) Rat trabeculaeTriton X-1001180- Luo et al. (2002) Rabbit skeletalTriton X-10011807.0 Fukuda et al. (2003) Bovine cardiacTriton X-10011807.0 Prado et al. (2005) Rabbit skeletalXTriton X-100-1807.0 Fukuda et al. (2005) Bovine and rat cardiacTriton X-10011807.0 Stelzer et al. (2006) Mouse cardiacSaponin + Triton X-10011807.0 Terui et al. (2010) Pig cardiacTriton X-10011807.0 Gillis and Klaiman (2011) Fish cardiacTriton X-10011707.0 Curtin et al. (2015) Rabbit skeletalXTriton X-10022007.1 Li et al. (2016) Rabbit skeletalTriton X-100-1807.0 Land et al. (2017) Human cardiacTriton X-10012007.1 Stehle (2017) Guinea pig cardiacTriton X-100-1707.0 Breithaupt et al. (2019) Rat cardiacGlycerol + Triton X-10012007.0 Giles et al. (2019) Mouse cardiacSaponin + Triton X-10011807.0 Azimi et al. (2020) Rat skeletalDissection1-7.1 Rebbeck et al. (2020) Human and rat skeletalDissection1-7.4 Palmer et al. (2020) Mouse cardiacTriton X-10012007.0Open in a separate windowA mark (X) in the Intact column indicates studies that directly compared measurements on both intact and skinned muscle (either performed within the same study or by considering previously published results). Var, variable.Sarcomere structureThe geometrical configuration and separation of the myofilaments regulate their interaction in the native system and hence their ability to generate tension. Under normal physiological conditions, the filament lattice structure is influenced by a complex balance of opposing forces, which include (Millman, 1998) electrostatic interactions between both thick and thin filaments (with charge being affected by pH and screened by the surrounding ionic strength), van der Waals forces, and entropic thermal forces, as well as Donnan osmotic force (whereby water enters the filament lattice to dilute counterions surrounding the charged filaments; Ilani, 2015). It is therefore unsurprising that this balance becomes disrupted upon removal of the sarcolemma.Muscle skinning broadly conserves the sarcomere assembly, but, as illustrated below, detailed quantitative features are altered at different scales. Microscopy and synchrotron x-ray measurements on skinned muscle report a modest increase in sarcomere length (∼3%), accompanied by a greater lateral expansion (up to twofold, depending on conditions), compared with intact cells. This is apparent in both skeletal (Matsubara and Elliott, 1972) and cardiac muscle (Irving et al., 2000; Roos and Brady, 1989). In both skinned and intact preparations, longitudinal stretching decreases the myofilament lattice spacing monotonically. This occurs more slowly in the skinned system, especially at large sarcomere lengths (Fig. 1; Irving et al., 2000). Despite their similar overall behavior, different physical effects are likely to operate in the two systems. The volume of intact cells is approximately conserved (Yagi et al., 2004), and therefore, stretching the cell decreases its cross-sectional area. As the sarcomere number remains constant, this increases the sarcomere density and hence stress generation (force per unit cross-sectional area). The constant-volume constraint is removed in skinned systems (Godt and Maughan, 1977; Irving et al., 2000; Matsubara and Elliott, 1972), which allows the structure to respond more visibly to other forces.Open in a separate windowFigure 1.Average myofilament spacing as a function of the sarcomere length in intact and relaxed skinned rat trabeculae, measured by x-ray diffraction. Adapted from Irving et al. (2000).The expansion of the myofilament spacing in skinned preparations can be reversed by increasing the osmotic pressure of the solution using dextran (Cazorla et al., 2001; Konhilas et al., 2002). However, this compressive effect does not by itself return the myofilaments fully to their intact physiological state (Konhilas et al., 2002). Recent x-ray diffraction experiments have identified an alteration of the detailed molecular structure of the thick filaments below physiological temperatures (Caremani et al., 2019, 2021). Although this effect is overlooked in many experiments, it may significantly affect cross-bridge kinetics.Skinning may also impact sarcomere morphology on larger scales. While measuring the effect of skinning on the sarcomere length in rat heart trabeculae using laser diffraction, Kentish et al. (1986) observed an increase in the diffraction intensity and a decrease in the dispersion of the first-order diffraction. Although this effect might result from the loss of intracellular scatterers (mitochondria, cytosolic proteins, etc.) upon skinning, the authors hypothesize that the skinning process might effectively enhance the homogenization of the sarcomere environment of the skinned tissue, relative to the intact one, where individual cells may display spontaneous and uncoordinated contractions. Nonetheless, the relative homogeneity of the skinned tissue degrades rapidly after successive contractions, possibly due to a loss of integrity of the cellular structure and content, in both cardiac (Kentish et al., 1986) and skeletal muscle (Fabiato and Fabiato, 1978b). This reflects a degree of irreproducibility inherent to skinned systems.Sarcomere structure strongly regulates contractile properties. Changes in both sarcomere length and interfilament spacing affect cross-bridge cycling and influence the regulation and amount of tension generated by skinned sarcomeres. Recent evidence also suggests that skinning may perturb myofilament interactions via steric effects due to myosin head orientations (Caremani et al., 2019, 2021; Konhilas et al., 2002). These effects, discussed further below, highlight the complexity in the disruption of the sarcomere function caused by skinning, relative to intact muscle, and the challenge in rationalizing their discrepancies based on fundamental physics principles. Ultimately, the extent to which skinning modifies sarcomere functionality bears critically on the interpretation of skinned muscle experiments.Passive mechanical compliancePassive mechanical properties of cardiac muscle strongly govern diastolic behavior. In intact tissue, these may have contributions originating in the cells themselves and the extracellular matrix (mostly comprising collagen). Passive tension and sarcomere length vary nonlinearly in both intact and skinned rat ventricular trabeculae preparations (Fig. 2; Kentish et al., 1986). However, in the skinned case, this length dependence is weaker, and the extension range is greater, indicating the presence of additional parallel elastic elements in the intact tissue, potentially associated with the sarcolemma or extracellular structures.Open in a separate windowFigure 2.Passive stress increasing with sarcomere length in skinned and intact rat ventricular trabeculae. The skinned results indicate enhanced mechanical compliance. Adapted from Kentish et al. (1986). Fig. 2 is reprinted with permission from Circulation Research.The qualitative similarity in the passive force-length relations in intact and skinned muscle makes the attribution of their quantitative differences challenging. The direct contribution of the sarcolemma itself, although plausible in principle, is expected to be weak, given its high compliance. However, it is more likely to contribute indirectly, given that the cell volume remains approximately constant upon stretching (Yagi et al., 2004). This effect may also be exacerbated by the Coulombic repulsion of the negatively charged myofilaments that, when confined within a fixed volume, would enhance resistance to lateral cellular compression (Kentish et al., 1986). Skinning may also cause the loss of intracellular components that contribute to the passive mechanics, e.g., a nonfilamentous stroma, comprising vesicular elements that dissolve in the skinning process (Kentish et al., 1986). Similarly, the loss of tubulin dimers from the cytoplasm may interfere with the viscoelastic behavior and resistance to cell shortening of the microtubule cytoskeleton (White, 2011).Structural differences can also explain discrepancies between skinned and intact muscle properties. Variations in the ionic strength acting on skinned myocytes have identified a mechanical contribution from the intracellular cytoskeleton (Roos and Brady, 1989). Similarly, titin contributes to the passive stiffness in isolated myofibrils and skinned single fibers, separately from the extracellular (mostly collagen) contribution (Cazorla et al., 2001; Fukuda and Granzier, 2005; Fukuda et al., 2005; Herzog, 2018; Powers et al., 2017). Within the isolated sarcomeric system, the stiffness varies inversely with the titin molecular size (Mijailovich et al., 2019; Prado et al., 2005), but this correlation disappears in intact fiber bundles, where extracellular contributions (e.g., from collagen) may dominate (Brower et al., 2006; Chung and Granzier, 2011; Fomovsky et al., 2010).Although the above observations highlight the limitations of using skinned preparations as a model for investigating passive mechanics in intact tissue, there may be indirect implications for contractile function. The distribution of force between passive and active mechanisms affects contraction, e.g., via force-dependent Ca²⁺ sensitivity (Cazorla et al., 2001; Fukuda and Granzier, 2005; Fukuda et al., 2005; Martyn and Gordon, 2001; Mijailovich et al., 2019; Sweitzer and Moss, 1990). In particular, passively elastic titin influences active contraction via the release of troponin I (TnI) from actin, as a result of the redistribution of mechanical load and strain on both the thick and thin filaments (Mijailovich et al., 2019). It may also determine the sarcomere length for a given afterload or the shortest sarcomere length in isotonic contractions.Calcium dependence of tension generationSkinned preparations are often used to measure the Ca²⁺ dependence of force development under equilibrium conditions. Measured F-pCa relations (e.g., Fig. 3) are conventionally characterized by their maximum saturating value, the location of the half-maximum point (the “sensitivity,” pCa₅₀), and the Hill coefficient n (quantifying the rate of rise and taken as a measure of cooperativity). To assess their validity, analogous F-pCa relations may also be generated in intact muscle by controlling the intracellular [Ca²⁺] homeostasis via tetanization, i.e., high-frequency activation (Fig. 3). Reported F-pCa relationships vary significantly according to the muscle type and preparations (Fabiato, 1981; Fukuda et al., 2003; Hibberd and Jewell, 1982; Kentish et al., 1986). This is problematic insofar as measurements in skinned systems aim to reproduce the “authentic” behavior in the intact system. The most intuitive mechanism involves an increased Ca²⁺-troponin binding affinity (Allen and Kentish, 1985; Kentish et al., 1986; Stephenson and Wendt, 1984), but more complex contributions also originate in the thick-filament structure upon stretching (Zhang et al., 2017).Open in a separate windowFigure 3.Comparing the force-calcium relationship in intact and skinned muscle. (a) Intact (ferret, 30°C; Yue et al., 1986) versus skinned (rabbit, 29°C; Harrison and Bers, 1989) muscle. (b) Pooled measurements derived from intact (solid symbols, pCa₅₀ ≈ 6.21, n ≈ 4.9) and skinned (open symbols, 6.04, 3.8) preparations of the same rat ventricular myocytes. max, maximum. From Gao et al. (1994). Fig. 3 is reprinted with permission from Circulation Research.Both pCa₅₀ and n are significantly enhanced in the intact case (in ferret) relative to skinned tissue (rabbit), substantially exceeding typical species-dependent variability observed in skinned muscle (Fig. 3 a; Bers, 2001). A similar qualitative conclusion was drawn from comparisons of intact and skinned preparations of the same rat ventricular myocytes (Fig. 3 b; Gao et al., 1994). These discrepancies are particularly significant when comparing the measured sensitivity values (pCa₅₀ = 5.52; Land et al., 2017) with physiological systolic [Ca²⁺] levels in the heart (0.6 µM ≃ pCa 6.22; Coppini et al., 2013; Land et al., 2017). Thus, the skinned muscle measurements are clearly incompatible with observed physiological behavior in intact myocytes and hence at the organ scale. Although the dominant underlying biophysical reason for these differences is uncertain, the detailed experimental conditions are fundamentally important (Bers, 2001). A rigorous quantitative comparison is therefore challenging.Skinning may affect the F-pCa relation via the sarcomere structure. An increase in the myofilament spacing plausibly reduces the rate of myosin cross-bridge formation and hence the amount of force generated for a given [Ca²⁺]. This would translate into a reduction in pCa₅₀, induced by muscle shortening, as observed in both skinned and (more weakly) intact preparations (Komukai and Kurihara, 1997). This mechanism may arguably contribute to the Frank–Starling mechanism in muscle, whereby the strength of contraction increases with stretch. However, this intuitive explanation has been shown to be insufficient in accounting for the complete effect on calcium sensitivity (Irving and Craig, 2019; de Tombe et al., 2010). It is also contradicted by experiments in which comparable myofilament spacings were achieved either via dextran-based osmotic compression or by sarcomere stretching (Konhilas et al., 2002). These discrepancies suggest that the filament spacing may not be the dominant contributor to pCa₅₀. However, this conclusion assumes the functional equivalence of the two scenarios. This may not be the case, as skinning may perturb other intracellular structures (e.g., titin or thin-filament regulatory proteins; Komukai and Kurihara, 1997). Experiments on mouse skinned cardiomyocytes have suggested that titin regulates filament spacing (Cazorla et al., 2001). Osmotic pressure may also impact the cross-bridge structural configuration on smaller molecular scales (Caremani et al., 2021; Konhilas et al., 2002).The sensitivity of the myofilaments to their chemical environment adds a further layer of complexity to skinned experiments. As discussed further below, F-pCa curves depend on the ionic strength, [Mg²⁺], and pH, all of which are routinely specified in skinned-experiment protocols. Skeletal muscle measurements have shown that increasing the temperature of the bathing solution increases the [Ca²⁺] required to activate skinned muscle as well as the maximal generated force (Godt and Lindley, 1982). Similarly, decreasing [Mg²⁺] lowers the activation [Ca²⁺] (Godt and Lindley, 1982). However, the native cell features other regulators that are lost during skinning and are not typically included in experiments. Sensitizers like taurine, carnosine-like compounds, and myosin light-chain kinase modestly increase the Ca²⁺ sensitivity (Gao et al., 1994). β-Adrenergic stimulation of intact muscle activates PKA, which in turn affects sarcomere dynamics by phosphorylating TnI and myosin-binding protein C (Gillis and Klaiman, 2011; Kentish et al., 2001; Patel et al., 2001). TnI phosphorylation decreases its binding affinity for Ca²⁺ (de Tombe and Stienen, 1995; Patel et al., 2001; Zhang et al., 1995), while that of myosin-binding protein C induces a movement of the myosin heads that accelerates force development.Despite their appealing relative simplicity, inconsistencies between skinned and intact muscle suggest fundamental alterations to muscle function by the skinning process. Following the rapid length release and restretch of skinned rat trabeculae, force redevelopment is Ca²⁺-dependent (Wolff et al., 1995b), unlike the rate of force redevelopment after a rapid-length release of intact ferret trabeculae (Hancock et al., 1993). This discrepancy is arguably explained by the relative dominance of thin- or thick-filament kinetics, respectively (Hunter et al., 1998).Taken together, these results illustrate the challenge of objectively determining the physiological Ca²⁺ dependence of muscle tension, in large part owing to the considerable technical challenge of replicating the native conditions of the myofilament system in vitro.Force-length relationThe sarcomere length dependence of force generation that underlies the Frank–Starling mechanism is a fundamental property of muscle behavior. Contributing mechanisms include the variation in myofilament overlap as the sarcomere is stretched, the apparent increase in the binding of Ca²⁺ to TnC with increasing length (Hibberd and Jewell, 1982; Kobirumaki-Shimozawa et al., 2014), and the modulation of the thick- (Fukuda et al., 2001; Zhang et al., 2017) and thin-filament structures (Zhang et al., 2017). The passive mechanical properties of titin (which vary according to the isoform) affect the variation in the lattice spacing under tension, and hence the length dependence of the actomyosin interaction (Fukuda et al., 2003). Recent evidence shows that the strain on titin, effectively acting as a force sensor, contributes to the Frank–Starling effect by influencing the structure of both the thin and thick filaments that are different from Ca²⁺-induced changes (Ait-Mou et al., 2016).Length-dependent tension, manifested in the F-pCa relationship, is qualitatively similar in intact and skinned preparations (Fig. 4). In the intact case, active tension was measured as the difference between the maximum tension in transiently stimulated muscle and the resting (unstimulated) tension at the same sarcomere lengths. The process was repeated at different [Ca²⁺] values in the bathing solution, so as to modulate the intracellular calcium. Comparing Fig. 4, a and b, for sufficiently low [Ca²⁺] below the level for full activation, the skinned- and unskinned-tissue measurements show a qualitatively similar transition from a concave to a convex dependence as [Ca²⁺] is increased. The results suggest that, whereas the unskinned system sustains no active tension for sarcomere lengths below ∼1.6 µm, the skinned preparation allows tension generation in this regimen, albeit at unphysiologically large [Ca²⁺]. However, the ability to measure (potentially heterogeneous) sarcomere lengths accurately in this regimen is questionable.Open in a separate windowFigure 4.Active force generation in intact and skinned rat ventricular trabeculae as a function of sarcomere length, for different bath [Ca²⁺]. From Kentish et al. (1986). Fig. 4 reprinted with permission from Circulation Research.For sufficiently low [Ca²⁺], the basic contraction mechanisms are thus preserved after skinning, at least qualitatively, suggesting that the general features of the force-length relationship are inherent myofibril properties. However, this conclusion assumes that (1) the chemical environments of the myofilaments are largely similar (any experimentally defined environment can only approximate the real cytosol), and (2) myofilament properties are not appreciably modified by the skinning process. The latter condition may be affected by the reported swelling of the myofilament lattice (Godt and Maughan, 1977; Irving et al., 2000; Konhilas et al., 2002; Matsubara and Elliott, 1972) or by any damage to the filaments occurring during the skinning process. Both of these effects should reduce the gradient of the tension relative to stretch.Significant variations in measurements may originate from structural causes at different levels. The above results, derived from trabeculae, show a steeper length dependence for short sarcomere lengths, compared with those of Fabiato and Fabiato (1975) on (mechanically) skinned maximally activated single ventricular myocytes (Kentish et al., 1986). This discrepancy might be ascribed either to the conservation of intercellular connections and extracellular connective tissue that might be lost in the skinned single myocytes, or to differences in the myofilament spacing in the multicellular tissue preparation. Some more subtle effects, such as the temperature-dependent alteration of the internal thick-filament structure in demembrenated muscle, observed recently (Caremani et al., 2019, 2021), seldom receive due consideration.Length-dependent F-pCa measurements show the sensitivity of muscle activation by calcium increasing with length, as marked by an increase in pCa₅₀ (Fig. 5). The maximum generated force at saturating [Ca²⁺] also increases. However, the Hill coefficient (n ≈ 7) does not vary significantly. A small but statistically significant increase in n was previously reported (Kentish et al., 1986), albeit based on sparser data, and was explained by invoking several mechanisms, e.g., interactions between adjacent tropomyosin molecules or alterations to the number of possible cross-bridges. Nonetheless, significant discrepancies even in the absolute values of n reported in other studies are also highlighted, potentially related to experimental conditions and the choice of skinning protocol.Open in a separate windowFigure 5.Dependence of the calcium sensitivity on sarcomere length. (a) Hill-type F-pCa for sarcomere lengths (SLs) = 1.85, 1.95, 2.05, 2.15, and 2.25 µm. Forces are normalized to the maximum force measured at SL = 2.05 µm. The data do not show a change in the Hill coefficient. (b) Increase in the Ca²⁺ sensitivity (decreasing [Ca²⁺] at half-maximum) with increasing SL, measured from the position of the inflection point in the fitted Hill curves from panel a. Adapted from Dobesh et al. (2002).The force-length relation in striated muscle underpins its central physiological role. Whereas the appeal of skinned muscle experiments for characterizing force generation is highlighted by numerous experiments, rationalizing quantitative differences remains notoriously challenging. In large part, this stems from the highly multifarious influence of the skinning process on the intracellular system and on details of the preparation protocol.Practical challenges: performing skinned muscle experimentsThe previous section illustrated the ability of skinned muscle preparations to reproduce intact muscle behavior while highlighting significant quantitative differences between the two systems. Clarifying the sources of these differences is crucial when developing practical applications that seek to exploit skinned muscle as a reductionist model for native-state muscle. One important hurdle is to correctly replicate the chemical and physiological intracellular environment, in particular with regard to [Mg²⁺], [ATP], pH, and the ionic strength. By tuning the experimental parameters to match the physiological conditions, the consistency between skinned and intact systems can be significantly improved (Gao et al., 1994; Mijailovich et al., 2021). Over decades, systematic efforts have sought to achieve this through detailed computations of the chemical equilibria of the bathing solutions (Fabiato, 1985a; Fabiato and Fabiato, 1975, 1977; Godt and Maughan, 1977; Moisescu, 1976). In practice, experimental protocols vary, sometimes idiosyncratically, between laboratories.This section outlines some of the elements of experimental protocols for skinned muscle that pose particular challenges insofar as they may significantly impact measurement outcomes.Bathing solution composition

ATP

After skinning, mitochondrial function is compromised, and hence, myocytes can no longer produce ATP (Rüegg, 2012). In multicellular tissue experiments, even a plentiful supply of ATP in the bathing solution may diffuse too slowly to maintain a homogeneous concentration throughout the fiber network (Godt, 1974). However, the inherent ATPase activity of muscle contraction implies a consumption of ATP supplies over the time of experiments. ATP-regenerating systems include creatine phosphate (typically 10–15 mM; Godt, 1974; Lamb and Stephenson, 2018). Nonetheless, in multicellular tissue, the rapid hydrolysis of ATP within the contractile system may yet produce an ATP concentration gradient between the interior and exterior of the network that inaccurately reflects the native state. This problem is arguably less serious in cardiac than skeletal myocytes (typical cardiac cell diameters are ∼13−20 µm, and lengths are ∼60−120 µm [Campbell et al., 1987, 1989; Liu et al., 1991], whereas skeletal muscle fiber diameters range from several microns to thousands of microns [Jimenez et al., 2013], with lengths sometimes reaching centimeters). However, the problem may yet arise in trabeculae.The physiological role of ATP in a given experiment, in addition to its participation in cross-bridge cycling, depends on the muscle preparation. In skeletal muscle experiments that preserve intracellular membrane structures (Endo and Iino, 1980; Launikonis and Stephenson, 1997), ATP governs calcium pumping into the SR (Godt, 1974; Lamb and Stephenson, 2018). This function is of course nonexistent in preparations where the SR has been dissolved. Alongside its role as energetic fuel, ATP also maintains the extensibility of the muscle by allowing myosin to dissociate from actin (Best et al., 1977; Weber and Murray, 1973).The decrease in maximum force with increasing [ATP] (in its physiological form MgATP; Fig. 6 b) is intuitively explained by the reduction in the number of formed cross-bridges (since ATP binding is associated with the release of rigor myosin; Best et al., 1977). An accompanying decrease in pCa₅₀ and an increase in the Hill coefficient (Fig. 6 a; Best et al., 1977) are both complicated by their Mg²⁺ dependence. These observations have been explained in terms of the effective cooperativity between neighboring cross-bridges in altering the inhibitory properties of troponin, which would arguably increase cross-bridge activation at a given [Ca²⁺] (Best, 1983; Best et al., 1977; Weber and Murray, 1973). However, this scenario is difficult to reconcile with analogous studies in skeletal muscle that report a qualitatively similar behavior for pCa₅₀ but with little [MgATP] dependence on maximum tension (Godt, 1974).Open in a separate windowFigure 6.Dependence of the force–calcium relationship on MgATP in the rat heart. (a) Decrease in Ca²⁺ sensitivity (increase in [Ca²⁺] at half-maximum) as [MgATP] increases from 30 to 100 µM ([Mg²⁺] = 50 µM). (b) Decrease in the maximum tension with increasing [MgATP]. Adapted from Best et al. (1977).

Mg²⁺

Mg²⁺, the second most abundant cation in muscle cells after K⁺, regulates the Ca²⁺ sensitivity of myofilament activity via its binding affinity to troponin (Alpert et al., 1979; Bers, 2001; Best, 1983; Best et al., 1977; Rayani et al., 2018; Tikunova and Davis, 2004). The Ca²⁺-specific low-affinity binding site (site II) at the N-terminal end of cardiac TnC serves as the principal initiator of contraction in the presence of Ca²⁺ (Bers, 2001). However, the structure of TnC is also controlled by binding sites III and IV, located at the C-terminal end, which competitively bind either Ca²⁺ (with high affinity) or Mg²⁺ (low affinity; Rayani et al., 2018; Tikunova and Davis, 2004). According to some cardiac muscle experiments, more Ca²⁺ is required to achieve a given degree of activation as [Mg²⁺] increases in the millimolar range (Best, 1983; Tikunova and Davis, 2004), consistent with competitive binding of these ions on TnC. However, this interpretation is contested by other cardiac experiments claiming negligible impact to the Ca²⁺ sensitivity under even an order-of-magnitude change in Mg²⁺ (Allen et al., 2000). The precise effect of Mg²⁺, while being potentially artifactual in some cases, may also vary with the dominant mechanism of action in the specific muscle system considered.Historically, setting the physiologically correct [Mg²⁺] has been challenging. Its determination requires the consideration of multiple binding equilibria and is naturally prone to uncertainty (Lamb and Stephenson, 2018). Given its relative abundance, cytosolic Mg²⁺ was initially assumed to merely ensure the balance for anionic charge, but its regulatory role was recognized subsequently. Various techniques have measured [Mg²⁺] (using spectrophotometry, Mg²⁺-sensitive electrodes, dye-based measurements, etc.). However, these measurements carry significant uncertainties, particularly given the difficulty of discerning free cytosolic Mg²⁺ from the total cellular magnesium (up to 20 times greater, contained in MgATP or cellular compartments) or interference from other ions (Romani and Scarpa, 1992). Many measurements report [Mg²⁺] as being consistently 0.4–0.8 mM but reaching up to 3.5 mM in some cases (Romani and Scarpa, 1992). In the intact rat heart specifically, values of 0.72 mM (from epifluorescence; Gao et al., 1994) or 0.85 mM (¹⁹F-NMR; Murphy et al., 1989) have been measured. [Mg²⁺] in excess of several millimolars are used in some studies but are known to be above the physiological level (Bers, 2001; Hunter et al., 1998).

pH

Intracellular pH in intact muscle regulates all the stages of tension generation, including the handling of Ca²⁺ by sarcolemmal electrophysiology, its delivery to the myofilaments, and the response of the filaments to the Ca²⁺ signal (Orchard and Kentish, 1990). This versatility makes it difficult to establish the relative significance of pH on sarcomere function specifically.In skinned muscle, a decrease in pH decreases pCa₅₀. The results in Fig. 7 show a 0.1% drop in pH producing a 0.1% drop in pCa₅₀ (Bers, 2001; Orchard and Kentish, 1990). The precise mechanism for this effect remains uncertain but may involve competition of H⁺ with Ca²⁺ for binding to TnC, interactions within the troponin complex, or the shielding of the net effective negative charge of the TnC binding site (Orchard and Kentish, 1990). Although a decrease in calcium sensitivity was also confirmed qualitatively in tetanized intact cardiac muscle (Marban and Kusuoka, 1987), the results differ quantitatively.Open in a separate windowFigure 7.Dependence of pH on the force-calcium relationship in guinea pig trabeculae. Adapted from Orchard and Kentish (1990).The observed decrease in maximal force resulting from decreasing pH in skinned muscle may be due to a direct impact on the efficiency of the coupling of ATP hydrolysis to cross-bridge force generation (Fig. 7; Orchard and Kentish, 1990). ATPase activity is affected by pH in intact muscle, albeit more weakly (Blanchard and Solaro, 1984; Kentish and Nayler, 1979; Orchard and Kentish, 1990). However, it is uncertain whether the same dominant mechanisms are relevant in the intact and skinned cases.The suitability of skinned muscle experiments for reliably investigating pH dependence is thus questionable. Bathing solutions for skinned muscle are typically designed with a high pH-buffering capacity (e.g., with 90 mM HEPES) to maintain a stable pH ∼7 (see Lamb and Stephenson, 2018).

Ionic strength

Ionic strength impacts inversely on the maximum force generated by skinned muscle (Fig. 8; Kentish, 1984). In practice, it can be controlled experimentally, in both cardiac and skeletal experiments, for example by varying KCl in the bathing soution (Kentish, 1984; Solaro et al., 1976). Reported ionic strength values range between 150 and 200 mM (Fig. 8). The inhibition of tension appears to be associated with Ca²⁺ binding, as this ionic strength dependence is [Ca²⁺] dependent only in the presence of MgATP (in skeletal muscle; Solaro et al., 1976). However, the precise ionic strength in intact muscle is uncertain (Gao et al., 1994), as reflected in the lack of consensus in the literature (see Open in a separate windowFigure 8.Dependence of generated tension on osmolarity. The osmolarity Γ/2 was controlled by varying (a) the Cl⁻ salt (filled circles: KCl; open circles: NaCl; diamonds: TMACl; triangles: choline Cl) or (b) K⁺ salt concentrations (filled circles: KCl, filled squares: K propionate; open square: K Mes), for pCa = 3.8. The consistency between the results suggests that the tension depends predominantly on the ionic strength rather than on the size of specific ions. From Kentish (1984). Fig. 8 reprinted with permission from Journal of Physiology.

Conclusion

The above considerations of ATP, Mg²⁺, pH, and ionic strength highlight the sensitivity of skinned muscle measurements to the precise solution composition. Establishing the correct recipe is made all the more challenging given that the impact on measured force generation varies between muscle systems and species. As argued above, although differences between measurements often appear to be quantitative, this does not exclude the possibility of qualitative differences in the dominant mechanisms of action. This fundamental ambiguity introduces considerable complication in translating results meaningfully to the intact system.TemperaturePhysiological function emerges from the balance of multiple temperature-dependent processes. Although measurements should thus ideally always be done at physiological temperature, lower temperatures are often used in practice due to the impaired stability of the sarcomere structure in skinned preparations at higher temperatures. This can have significant consequences on contraction, given the highly variable temperature sensitivities of different subcellular mechanisms (Rall and Woledge, 1990).There is widespread agreement that cooling reduces the maximum generated force in a wide range of muscle types and preparations (Fig. 9; Fabiato, 1985b; Godt and Lindley, 1982; Harrison and Bers, 1989; Stephenson and Williams, 1985; Sweitzer and Moss, 1990). This result has been argued to result more from a change in the force exerted by cross-bridges than from the number of cross-bridges formed (Sweitzer and Moss, 1990). In contrast, the temperature dependence of calcium sensitivity is less consistent. Skinned muscle displays either an increase (Brandt and Hibberd, 1976; Harrison and Bers, 1989; Orentlicher et al., 1977; Sweitzer and Moss, 1990) or a decrease in pCa₅₀ (Fabiato, 1985b; Godt and Lindley, 1982; Stephenson and Williams, 1985) with increasing temperature. However, the former result may be an artifact associated with heterogeneous shortening of sarcomeres at higher temperatures (Sweitzer and Moss, 1990).Open in a separate windowFigure 9.Temperature dependence of the F-pCa relationship in skinned trabeculae from the rabbit ventricle, showing an increase in both the maximum tension C_max and the sensitivity pCa₅₀ (pCa at half-maximum) with increasing temperature. Adapted from Harrison and Bers (1989).More recent work has revealed further complications in the regulatory role of temperature in muscle. In particular, temperature influences structural thick-filament regulation in both cardiac and skeletal muscle (Caremani et al., 2019, 2021; Park-Holohan et al., 2021). Reducing the temperature disrupts the orderly configuration of the myosin lever arms along the thick filaments, making them less available for force generation and causing an almost threefold decrease in total tissue force.The above experimental results highlight the multifaceted complexity of temperature dependence that arises from the interdependence of multiple molecular processes. Skinned preparations constitute only a subsystem within the overall muscle system, and there is therefore no guarantee that the kinetic balance within the reduced system is physiologically accurate.Sarcomere heterogeneityFor conceptual convenience, muscle tissue is often represented as a homogeneous assembly of identical sarcomeres acting in synchrony. This picture is simplistic in reality. Aspects of muscle dynamics, even under isometric conditions, derive specifically from the heterogeneous behavior at the sarcomere level. For example, within a myofibril, tension relaxation proceeds with the onset of rapid lengthening (“give”), initially in a single weak sarcomere, that then propagates to other sarcomeres along the myofibril (Edman and Flitney, 1982; Poggesi et al., 2005; Stehle, 2017). This effect accounts for the [P_i]-dependent asymmetry in the force kinetics that is observed in contraction-relaxation cycles when [Ca²⁺] is stepped up and down (Poggesi et al., 2005). It also suggests that relaxation kinetics is governed not only by the rate-limiting steps of the cross-bridge cycle of a generic myosin molecule but also by collective effects at a higher structural level.This effect arguably escapes notice in skinned-fiber experiments that exploit the flash photolysis of caged compounds to time-resolve the details of cross-bridge–cycle kinetics (e.g., the photorelease of inorganic phosphate P_i modulates cross-bridge kinetics; Araujo and Walker, 1996; Dantzig et al., 1992; Millar and Homsher, 1990; Tesi et al., 2000). These experiments suffer from important practical limitations. In particular, the relatively modest (unidirectional) changes in [P_i] achievable by photorelease fail to disrupt the chemomechanical equilibrium of the sarcomeres sufficiently to generate heterogeneous give. Under these near-equilibrium conditions, observed changes in force are more likely to reflect rate-limiting single-cross-bridge kinetics than transients in sarcomere heterogeneity. This obstacle was bypassed in experiments done on isolated myofibrils, which, in contrast, allow sufficiently large jumps in [P_i] (in both directions) to be imposed by rapid solution change (Poggesi et al., 2005; Stehle, 2017). By monitoring the progression of tension decay in conjunction with the lengths of individual sarcomeres, these experiments highlight the role of sarcomere dynamics in accounting for tension relaxation. Compared with skinned-tissue experiments, they also provide better consistency with the relaxation kinetics (k_TR) observed in mechanically induced force redevelopment (Stehle, 2017).Practical considerationsThe preceding discussion has highlighted the value of skinned muscle in emulating the essential features of intact muscle contraction in vivo. On the other hand, we have also described how discrepancies between intact and skinned muscle properties are sufficiently significant as to mar the prospect of considering skinned preparations as unambiguous surrogates. The underlying causes are complex, and it is often difficult to distinguish between experimental artifacts and manifestations of genuine physiological differences. This complexity is further compounded by species- or system-dependent specificities (e.g., cardiac versus skeletal muscle). Consequently, in practice, experimental protocols often evolve organically within laboratory communities, based on direct observations and acquired practical knowhow. Interestingly, a recent meta-analysis of published measurements of specific force in skinned human skeletal muscle noted a greater consistency in the results obtained within research groups (defined in terms of commonalities in authorship) than between them (Kalakoutis et al., 2021). This observation could be interpreted as revealing a genealogy of sorts in the evolution of protocols that is at odds with rigorous and objective development, thereby possibly mitigating the appeal of the experiments altogether.Tempting as it may be to imagine a universally applicable method, we feel it would be counterproductive to seek to disentangle and confront the rationales of individual protocols, with the risk of dogmatically promoting one valid method among several. The very idea of a unique universal recipe, valid for all experiments, is indeed highly questionable. As a more fruitful approach, we instead present the following themes as set of general guiding principles for encouraging good experimental practice.Monitoring sarcomeric dynamicsGiven the importance of sarcomere length and interfilament dynamics in force generation, we recommend that mechanical force measurements be accompanied by the simultaneous measurement of striation patterns. This would include the mean sarcomere length and, ideally, an index of heterogeneity and/or stability. We recognize that these measurements may be particularly challenging in cardiac trabeculae.Fixing the pHEnsuring the constancy of pH is paramount for ensuring consistency in measurements. This is achieved by applying a suitable buffer, in many cases imidazole.Saturation with ATPA useful simplification of the experimental system is to ensure that the cross-bridge cycling kinetics is not rate-limited by ATP. In most cases, this can be achieved by using solutions with at least 4 mM free ATP.Careful control of [Ca²⁺]The importance of correctly determining the concentration of free Ca²⁺ cannot be sufficiently emphasized. Some laboratories use pCa solutions based on recipes that originate with Fabiato and Fabiato (1979) or Godt and Lindley (1982). Those wishing to make new recipes can consider using the MaxChelator software suite (Bers et al., 2010; Patton et al., 2004), which can provide appropriate stoichiometric concentrations of Ca²⁺, Mg²⁺, EGTA, and ATP for use in experimental solutions. A useful recipe for producing buffers with varying [Ca²⁺] is to prepare “low” and “high” reference buffers (e.g., with pCa = 9.0 and 4.5) and to mix them in appropriate proportions.Choice of temperatureGiven the importance of temperature as a determinant of muscle kinetics, it stands to reason that experiments should be done at physiological temperatures. However, a practical drawback is its destabilization of the sarcomere structure. Skeletal fibers have historically been measured at lower temperatures (sometimes even near above freezing) to ensure that preparations last the experiment duration. Many experiments on both skeletal and cardiac muscle can be done at 15°C. However, it is worth noting that rodent myocardium is more fragile than human (where room temperature or even 37°C is possible), possibly owing to differences in metabolic and ATPase rates. As a general recommendation, we would encourage experimentalists to choose temperatures that are nearest to physiological conditions where the preparation is stable. It is, however, perhaps even more important to only compare experimental results obtained at the same temperature.ConclusionThe aim of this review was to survey the benefits of skinned muscle measurements for characterizing cardiac muscle physiology, while highlighting intrinsic challenges for both the conduct and the interpretation of measurements. These features are summarized in Strengths• Direct access to the sarcomere system• Separation of cellular subsystems (e.g., sarcomeres versus sarcolemma)• Ability to use fluorescent probes and other analytic tools• Convenience of controllably performing different standardized experiments (e.g., isometric/isotonic contractions)• Ability to perform protein exchange experiments that preserve overall functionality (e.g., troponin; Babu et al., 1988; Brenner et al., 1999; Gulati and Babu, 1989); and to probe time-resolve sarcomere dynamics by 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])• Simpler handling and storage logistics (samples can be thawed and analyzed after prior freezing) Weaknesses • Challenge of reproducing the native physiological environment• Variations in results between laboratories• Instability and sensitivity to temperature• Challenges of [Ca²⁺] calibration• Structural changes caused by skinning (e.g., altered sarcomere morphology, loss of cellular heterogeneity), impacting functional behaviorOpen in a separate windowThe potential pitfalls of mischaracterizing sarcomere behavior, based on skinned muscle measurements, are particularly exposed when considering the broader physiological context, where different cardiac subsystems operate simultaneously (Mosqueira et al., 2019; Niederer et al., 2019b). Pharmacological research increasingly exploits skinned muscle experiments to assess targeted drug action on sarcomeres (Dou et al., 2007; Edes et al., 1995; Fitton and Brogden, 1994; Hara et al., 1999; Kobayashi et al., 1991; Lamont and Miller, 1992; Lee and Allen, 1997; Lues et al., 1988; Scheld et al., 1989; Solaro and Rüegg, 1982; Sudo et al., 2001; Tadano et al., 2010). However, drug impact is notoriously multifaceted, and side effects, unseen in the isolated sarcomeres, may readily and unpredictably overwhelm intended effects (Lee and Allen, 1997; Lues et al., 1993). These side effects notwithstanding, the extrapolation of skinned-muscle measurements to the native cellular state and to systemic cardiac function encounters significant interpretational hurdles, as illustrated above.Skinned muscle measurements carry intrinsic uncertainty, as experiments performed using different animal models, temperatures, and protocols occasionally produce contradictory characterizations. Approximate quantitative accuracy is obviously highly problematic in the perspective of developing customized clinical care. This requirement is particularly important given the modular nature of models and the need to combine interacting subsystems on different length scales (Niederer et al., 2019a, 2019b). In practice, the interfacing of such modules normally requires ad hoc empirical alterations to model parameters, often relying on the modeler’s judgment (Hunter et al., 1998; Land et al., 2017). These choices are naturally often speculative.Despite these difficulties, it would be wrong to misrepresent the true potential of skinned-muscle experiments. Just as animal models are essential for investigating human physiology, skinned muscle provides an experimental setting with unique benefits. Biophysical modeling helps to formalize the conceptual basis for interpreting experimental data in terms of specific mechanisms (for example, an observed variation in pCa₅₀ may result from changes to troponin binding kinetics or cross-bridge formation). Global sensitivity analyses allow a ranking of the relative importance of individual model parameters, thus providing a handle for guiding judgment in how to use measurement-derived parameters (Longobardi et al., 2020). In this perspective, the benefit of models is in providing a framework for formulating and testing hypotheses, rather than delivering fixed and absolute representations of the muscle system.The appeal of skinned muscle preparations is best appreciated by seeing them not as a direct emulation of real muscle, but rather as one further element in the physiologist’s experimental armory. This issue is well illustrated by Irving and Craig (2019) with reference to a loosening of the thick-filament structure induced by cardiac myosin-binding protein C phosphorylation. This effect was manifested as a structural change in skinned cardiac muscle but may be eclipsed in the compact and crowded conditions of intact muscle. In such circumstances, attempting to reconcile the experiments, even qualitatively, may seem futile. Yet the skinned-muscle effect may well be the telltale indicator of a genuine regulatory mechanism that would otherwise remain invisible and unmeasurable in the intact system. Rather than seeking a literal mirroring of these skinned and intact experiments at any cost, additional physiological insight might potentially be gained by further pursuing the experiments, and comparing their quantitative results in parallel, in other cell types or under different experimental conditions. Ultimately, the integration of experimental findings remains a continual process involving a balance of pragmaticism and biophysically guided scientific judgment.     

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     

Imaging cell biology in live animals: Ready for prime time

Time-lapse fluorescence microscopy is one of the main tools used to image subcellular structures in living cells. Yet for decades it has been applied primarily to in vitro model systems. Thanks to the most recent advancements in intravital microscopy, this approach has finally been extended to live rodents. This represents a major breakthrough that will provide unprecedented new opportunities to study mammalian cell biology in vivo and has already provided new insight in the fields of neurobiology, immunology, and cancer biology.The discovery of GFP combined with the ability to engineer its expression in living cells has revolutionized mammalian cell biology (Chalfie et al., 1994). Since its introduction, several light microscopy–based techniques have become invaluable tools to investigate intracellular events (Lippincott-Schwartz, 2011). Among them are: time-lapse confocal microscopy, which has been instrumental in studying the dynamics of cellular and subcellular processes (Hirschberg et al., 1998; Jakobs, 2006; Cardarelli and Gratton, 2010); FRAP, which has enabled determining various biophysical properties of proteins in living cells (Berkovich et al., 2011); and fluorescence resonance energy transfer (FRET), which has been used to probe for protein–protein interactions and the local activation of specific signaling pathways (Balla, 2009). The continuous search for improvements in temporal and spatial resolution has led to the development of more sophisticated technologies, such as spinning disk microscopy, which allows the resolution of fast cellular events that occur on the order of milliseconds (Nakano, 2002); total internal reflection microscopy (TIRF), which enables imaging events in close proximity (100 nm) to the plasma membrane (Cocucci et al., 2012); and super-resolution microscopy (SIM, PALM, and STORM), which captures images with resolution higher than the diffraction limit of light (Lippincott-Schwartz, 2011).Most of these techniques have been primarily applied to in vitro model systems, such as cells grown on solid substrates or in 3D matrices, explanted embryos, and organ cultures. These systems, which are relatively easy to maintain and to manipulate either pharmacologically or genetically, have been instrumental in providing fundamental information about cellular events down to the molecular level. However, they often fail to reconstitute the complex architecture and physiology of multicellular tissues in vivo. Indeed, in a live organism, cells exhibit a 3D organization, interact with different cell types, and are constantly exposed to a multitude of signals originated from the vasculature, the central nervous system, and the extracellular environment. For this reason, scientists have been attracted by the possibility of imaging biological processes in live multicellular organisms (i.e., intravital microscopy [IVM]). The first attempt in this direction was in 1839, when Rudolph Wagner described the interaction of leukocytes with the walls of blood vessels in the webbed feet of a live frog by using bright-field transillumination (Wagner, 1839). Since then, this approach has been used for over a century to study vascular biology in thin areas of surgically exposed organs (Irwin and MacDonald, 1953; Zweifach, 1954) or by implanting optical windows in the skin or the ears (Clark and Clark, 1932). In addition, cell migration has also been investigated using transparent tissues, such as the fin of the teleost (Wood and Thorogood, 1984; Thorogood and Wood, 1987). The introduction of epifluorescence microscopy has enabled following in more detail the dynamics of individual cells in circulation (Nuttall, 1987), in tumors (MacDonald et al., 1992), or in the immune system (von Andrian, 1996), and the spatial resolution has been significantly improved by the use of confocal microscopy, which has made it possible to collect serial optical sections from a given specimen (Villringer et al., 1989; O’Rourke and Fraser, 1990; Jester et al., 1991). However, these techniques can resolve structures only within a few micrometers from the surface of optically opaque tissues (Masedunskas et al., 2012a). It was only in the early nineteen nineties, with the development of multiphoton microscopy, that deep tissue imaging has become possible (Denk et al., 1990; Zipfel et al., 2003b), significantly contributing to several fields, including neurobiology, immunology, and cancer biology (Fig. 1; Svoboda and Yasuda, 2006; Amornphimoltham et al., 2011; Beerling et al., 2011). In the last few years, the development of strategies to minimize the motion artifacts caused by the heartbeat and respiration has made it possible to successfully image subcellular structures with spatial and temporal resolutions comparable to those achieved in in vitro model systems, thus providing the opportunity to study cell biology in live mammalian tissues (Fig. 1; Weigert et al., 2010; Pittet and Weissleder, 2011).Open in a separate windowFigure 1.Spatial resolution and current applications of intravital microscopy. IVM provides the opportunity to image several biological processes in live animals at different levels of resolution. Low-magnification objectives (5–10×) enable visualizing tissues and their components under physiological conditions and measuring their response under pathological conditions. Particularly, the dynamics of the vasculature have been one of topic most extensively studied by IVM. Objectives with higher magnification (20–30×) have enabled imaging the behavior of individual cell over long periods of time. This has led to major breakthroughs in fields such as neurobiology, immunology, cancer biology, and stem cell research. Finally, the recent developments of strategies to minimize the motion artifacts caused by the heartbeat and respiration combined with high power lenses (60–100×) have opened the door to image subcellular structures and to study cell biology in live animals.The aim of this review is to highlight the power of IVM in addressing cell biological questions that cannot be otherwise answered in vitro, due to the intrinsic limitations of reductionist models, or by other more classical approaches. Furthermore, we discuss limitations and areas for improvement of this imaging technique, hoping to provide cell biologists with the basis to assess whether IVM is the appropriate choice to address their scientific questions.

Imaging techniques currently used to perform intravital microscopy

Confocal and two-photon microscopy are the most widely used techniques to perform IVM. Confocal microscopy, which is based on single photon excitation, is a well-established technique (Fig. 2 A) that has been extensively discussed elsewhere (Wilson, 2002); hence we will only briefly describe some of the main features of two-photon microscopy and other nonlinear optical techniques.Open in a separate windowFigure 2.Fluorescent light microscopy imaging techniques used for intravital microscopy. (A) Confocal microscopy. (top) In confocal microscopy, a fluorophore absorbs a single photon with a wavelength in the UV-visible range of the spectrum (blue arrow). After a vibrational relaxation (orange curved arrow), a photon with a wavelength shifted toward the red is emitted (green arrow). (center) In thick tissue, excitation and emission occur in a relative large volume around the focal plane (F.P.). The off-focus emissions are eliminated through a pinhole, and the signal from the focal plane is detected via a photomultiplier (PMT). Confocal microscopy enables imaging at a maximal depth to 80–100 µm. (bottom) Confocal z stack of the tongue of a mouse expressing the membrane marker m-GFP (green) in the K14-positive basal epithelial layer, and the membrane marker mTomato in the endothelium (red). The xy view shows a maximal projection of 40 z slices acquired every 2.5 µm, whereas the xz view shows a lateral view of the stack. In blue are the nuclei labeled by a systemic injection of Hoechst. Excitation wavelengths: 450 nm, 488 nm, and 562 nm. (B) Two- and three-photon microscopy. (top) In this process a fluorophore absorbs almost simultaneously two or three photons that have half (red arrow) or a third (dark red arrow) of the energy required for its excitation with a single photon. Two- or three-photon excitations typically require near-IR or IR light (from 690 to 1,600 nm). (center) Emission and excitation occur only at the focal plane in a restricted volume (1.5 fl), and for this reason a pinhole is not required. Two- and three-photon microscopy enable imaging routinely at a maximal depth of 300–500 µm. (bottom) Two-photon z stack of an area adjacent to that imaged in A. xy view shows a maximal projection of 70 slices acquired every 5 µm. xz view shows a lateral view of the stack. Excitation wavelength: 840 nm. (C) SHG and THG. (top) In SHG and THG, photons interact with the specimen and combine to form new photons that are emitted with twice or three times their initial energy without any energy loss. (center) These processes have similar features to those described for two- and three-photon microscopy and enable imaging at a maximal depth of 200–400 µm. (bottom) z stack of a rat heart excited by two-photon microscopy (740 nm) to reveal the parenchyma (green), and SHG (930 nm) to reveal collagen fibers (red). xy shows a maximal projection of 20 slices acquired every 5 µm. xz view shows a lateral view of the stack. Bars: (xy views) 40 µm; (xz views) 50 µm.The first two-photon microscope (Denk et al., 1990) was based on the principle of two-photon excitation postulated by Maria Göppert-Mayer in her PhD thesis (Göppert-Mayer, 1931). In this process a fluorophore is excited by the simultaneous absorption of two photons with wavelengths in the near-infrared (IR) or IR spectrum (from 690 to 1,600 nm; Fig. 2 B). Two-photon excitation requires high-intensity light that is provided by lasers generating very short pulses (in the femtosecond range) and is focused on the excitation spot by high numerical aperture lenses (Zipfel et al., 2003b). There are three main advantages in using two-photon excitation for IVM. First, IR light has a deeper tissue penetration than UV or visible light (Theer and Denk, 2006). Indeed, two-photon microscopy can resolve structures up to a depth of 300–500 µm in most of the tissues (Fig. 2 B), and up to 1.5 mm in the brain (Theer et al., 2003; Masedunskas et al., 2012a), whereas confocal microscopy is limited to 80–100 µm (Fig. 2 A). Second, the excitation is restricted to a very small volume (1.5 fl; Fig. 2 B). This implies that in two-photon microscopy there is no need to eliminate off-focus signals, and that under the appropriate conditions photobleaching and phototoxicity are negligible (Zipfel et al., 2003b). However, confocal microscopy induces out-of-focus photodamage, and thus is less suited for long-term imaging. Third, selected endogenous molecules can be excited, thus providing the contrast to visualize specific biological structures without the need for exogenous labeling (Zipfel et al., 2003a). Some of these molecules can also be excited by confocal microscopy using UV light, although with the risk of inducing photodamage.More recently, other nonlinear optical techniques have been used for IVM, and among them are three-photon excitation, and second and third harmonic generation (SHG and THG; Campagnola and Loew, 2003; Zipfel et al., 2003b; Oheim et al., 2006). Three-photon excitation follows the same principle as two-photon (Fig. 2 B), and can reveal endogenous molecules such as serotonin and melatonin (Zipfel et al., 2003a; Ritsma et al., 2013). In SHG and THG, photons interact with the specimen and combine to form new photons that are emitted with two or three times their initial energy (Fig. 2 C). SHG reveals collagen (Fig. 2 C) and myosin fibers (Campagnola and Loew, 2003), whereas THG reveals lipid droplets and myelin fibers (Débarre et al., 2006; Weigelin et al., 2012). Recently, two other techniques have been used for IVM: coherent anti-Stokes Raman scattering (CARS) and fluorescence lifetime imaging (FLIM). CARS that is based on two laser beams combined to match the energy gap between two vibrational levels of the molecule of interest, has been used to image lipids and myelin fibers (Müller and Zumbusch, 2007; Fu et al., 2008; Le et al., 2010). FLIM, which measures the lifetime that a molecule spends in the excited state, provides quantitative information on cellular parameters such as pH, oxygen levels, ion concentration, and the metabolic state of various biomolecules (Levitt et al., 2009; Provenzano et al., 2009; Bakker et al., 2012).We want to emphasize that two-photon microscopy and the other nonlinear techniques are the obligatory choice when the imaging area is located deep inside the tissue, endogenous molecules have to be imaged, or long-term imaging with frequent sampling is required. However, confocal microscopy is more suited to resolve structures in the micrometer range, because of the possibility of modulating the optical slice (Masedunskas et al., 2012a).

IVM to investigate biological processes at the tissue and the single cell level

The main strength of IVM is to provide information on the dynamics of biological processes that otherwise cannot be reconstituted in vitro or ex vivo. Indeed, IVM has been instrumental in studying several aspects of tissue physiopathology (Fig. 3, A and B). Although other approaches such as classical immunohistochemistry, electron microscopy, and indirect immunofluorescence may provide detailed structural and quantitative information on blood vessels, IVM enables measuring events such as variations of blood flow at the level of the capillaries or local changes in blood vessel permeability. These data have been instrumental in understanding the mechanisms of ischemic diseases and tumor progression, and in designing effective anticancer treatments.

Table 1.

IVM to study tissue physiopathology

The Golgi and the centrosome: building a functional partnership

The mammalian Golgi apparatus is characterized by a ribbon-like organization adjacent to the centrosome during interphase and extensive fragmentation and dispersal away from the centrosome during mitosis. It is not clear whether this dynamic association between the Golgi and centrosome is of functional significance. We discuss recent findings indicating that the Golgi–centrosome relationship may be important for directional protein transport and centrosome positioning, which are both required for cell polarization. We also summarize our current knowledge of the link between Golgi organization and cell cycle progression.

Introduction

The Golgi apparatus plays a central role in the secretory pathway. Newly synthesized proteins are transported from the ER to the Golgi, where they are posttranslationally modified. They are sorted into carriers for delivery to the plasma membrane or the endosomal–lysosomal system. The basic structural unit of the Golgi apparatus is a stack of flattened cisternae that is morphologically conserved among most species. In mammalian cells, individual Golgi stacks are connected laterally to form a continuous membranous system called the Golgi ribbon, which is located in close physical proximity to the centrosome (Fig. 1, left).Open in a separate windowFigure 1.The spatial relationship between the Golgi and the centrosome during the mammalian cell cycle. Golgi (red, stained with antibodies to GM130) and centrosome (green, stained with antibodies to centrin) staining of nonsynchronized bone cancer cells (U2-OS) shows the physical proximity between these two organelles during interphase (left) and its temporary loss during mitosis (right). Bar, 10 µm.The centrosome functions as the major microtubule-organizing center of the cell and plays an important role in cell polarization and ciliogenesis (Bettencourt-Dias and Glover, 2007). In a newly formed daughter cell, this nonmembrane-bound organelle is composed of a pair of centrioles that is surrounded by a cloud of electron-dense material called the pericentriolar matrix. γ-Tubulin ring complexes (γ-TuRCs) in the pericentriolar matrix allow the centrosome to nucleate the radial array of interphase microtubules whose minus ends are embedded in the centrosome and whose plus ends extend toward the cell periphery. After centrosome duplication in S phase, the two centrosomes move to opposite poles of the cell and become the spindle poles from which spindle microtubules grow. Centrosomes are generally located in the cell center close to the nucleus, although this central position is lost in response to a polarization stimulus, which prompts centrosomes to reorient toward the leading edge of the cell (Pouthas et al., 2008). In most cell types, centrosome reorientation is critical for the ability of cells to polarize and migrate (Yvon et al., 2002). The centrosome is also linked to ciliogenesis because one of its centrioles is converted into the basal body from which a primary cilium extends (D''Angelo and Franco, 2009).The spatial relationship between the Golgi apparatus and the centrosome is altered by changes in Golgi organization that occur during the cell cycle (Fig. 1). These two organelles are only adjacent in interphase when the Golgi apparatus is arranged as a ribbon in the pericentriolar region (Colanzi et al., 2003). In contrast, Golgi membranes are fragmented and dispersed throughout the cytoplasm during mitosis. Intriguingly, the pericentriolar localization of the Golgi is a feature typical of some eukaryotic cells, ranging from mammalian and amphibian cells (Thyberg and Moskalewski, 1999; Reilein et al., 2003) to amoeba (Rehberg et al., 2005). However, other eukaryotes, including plants and flies, have isolated Golgi stacks (Stanley et al., 1997; Nebenführ and Staehelin, 2001) or isolated cisternae in the case of Saccharomyces cerevisiae that are scattered throughout the cytoplasm without an obvious connection with the centrosome (Preuss et al., 1992).In this paper, we review recent findings indicating that the relationship between the Golgi and the centrosome in interphase is important for cell polarization. We also summarize the current understanding of how Golgi–centrosome interactions during mitosis affect cell division.

Are there functional interactions between the Golgi and the centrosome during interphase?

Golgi membranes are actively positioned in the pericentriolar position.

The localization of the mammalian Golgi ribbon next to the centrosome requires the microtubule and actin cytoskeleton (Brownhill et al., 2009). Microtubules have a dual role in organizing the pericentriolar Golgi ribbon. First, the subset of microtubules that is nucleated at the Golgi is necessary for the assembly of Golgi fragments into a connected ribbon in the cell periphery (Miller et al., 2009). Second, centrosomal microtubules provide the tracks along which Golgi membranes are transported to the cell center (Cole et al., 1996). Both steps depend on the minus end–directed motor complex dynein (Burkhardt et al., 1997; Miller et al., 2009). The actin cytoskeleton is also involved in localizing Golgi membranes. Actin fibers, which have been detected at the Golgi complex, are required for the maintenance of the pericentriolar position of this organelle by providing tracks for actin-based motors (Valderrama et al., 1998; Sahlender et al., 2005; Vicente-Manzanares et al., 2007). Actin fibers and microtubules are coordinated by proteins that associate with both cytoskeletal elements such as WHAMM, a Golgi-bound actin-nucleating factor, and MACF1, a microtubule–actin cross-linking protein (Lin et al., 2005; Campellone et al., 2008).Golgi organization and localization in the pericentriolar region also depend on Golgi-associated proteins (Ramirez and Lowe, 2009). Their depletion produces defects in Golgi organization ranging from a disconnected Golgi ribbon in the pericentriolar region (Puthenveedu et al., 2006) to dispersed ministacks in the cytoplasm (Diao et al., 2003; Yadav et al., 2009).

Table I.

Golgi-associated proteins that control the pericentriolar position of the Golgi apparatus

Mouse Models of Osteoarthritis: A Summary of Models and Outcomes Assessment

Osteoarthritis (OA) is a multidimensional health problem and a common chronic disease. It has a substantial impact on patient quality of life and is a common cause of pain and mobility issues in older adults. The functional limitations, lack of curative treatments, and cost to society all demonstrate the need for translational and clinical research. The use of OA models in mice is important for achieving a better understanding of the disease. Models with clinical relevance are needed to achieve 2 main goals: to assess the impact of the OA disease (pain and function) and to study the efficacy of potential treatments. However, few OA models include practical strategies for functional assessment of the mice. OA signs in mice incorporate complex interrelations between pain and dysfunction. The current review provides a comprehensive compilation of mouse models of OA and animal evaluations that include static and dynamic clinical assessment of the mice, merging evaluation of pain and function by using automatic and noninvasive techniques. These new techniques allow simultaneous recording of spontaneous activity from thousands of home cages and also monitor environment conditions. Technologies such as videography and computational approaches can also be used to improve pain assessment in rodents but these new tools must first be validated experimentally. An example of a new tool is the digital ventilated cage, which is an automated home-cage monitor that records spontaneous activity in the cages.

Osteoarthritis (OA) is a multidimensional health problem and a common chronic disease.³⁶ Functional limitations, the absence of curative treatments, and the considerable cost to society result in a substantial impact on quality of life.⁷⁶ Historically, OA has been described as whole joint and whole peri-articular diseases and as a systemic comorbidity.^9,111 OA consists of a disruption of articular joint cartilage homeostasis leading to a catabolic pathway characterized by chondrocyte degeneration and destruction of the extracellular matrix (ECM). Low-grade chronic systemic inflammation is also actively involved in the process.^42,92 In clinical practice, mechanical pain, often accompanied by a functional decline, is the main reason for consultations. Recommendations to patients provide guidance for OA management.^{22, 33,49,86} Evidence-based consensus has led to a variety of pharmacologic and nonpharmacologic modalities that are intended to guide health care providers in managing symptomatic patients. Animal-based research is of tremendous importance for the study of early diagnosis and treatment, which are crucial to prevent the disease progression and provide better care to patients.The purpose of animal-based OA research is 2-fold: to assess the impact of the OA disease (pain and function) and to study the efficacy of a potential treatment.^18,67 OA model species include large animals such as the horse, goat, sheep, and dog, whose size and anatomy are expected to better reflect human joint conditions. However, small animals such as guinea pig, rabbit, mouse, and rat represent 77% of the species used.^1,87 In recent years, mice have become the most commonly used model for studying OA. Mice have several advantageous characteristics: a short development and life span, easy and low-cost breeding and maintenance, easy handling, small joints that allow histologic analysis of the whole joint,³² and the availability of genetically modified lines.¹⁰⁸ Standardized housing, genetically defined strains and SPF animals reduce the genetic and interindividual acquired variability. Mice are considered the best vertebrate model in terms of monitoring and controlling environmental conditions.^7,14,15,87 Mouse skeletal maturation is reached at 10 wk, which theoretically constitutes the minimal age at which mice should be entered into an OA study.^64,87,102 However, many studies violate this limit by testing mice at 8 wk of age.Available models for OA include the following (32,111 physical activity and exercise induced OA; noninvasive mechanical loading (repetitive mild loading and single-impact injury); and surgically induced (meniscectomy models or anterior cruciate ligament transection). The specific model used would be based on the goal of the study.⁷ For example, OA pathophysiology, OA progression, and OA therapies studies could use spontaneous, genetic, surgical, or noninvasive models. In addition, pain studies could use chemical models. Lastly, post-traumatic studies would use surgical or noninvasive models; the most frequently used method is currently destabilization of the medial meniscus,³² which involves transection of the medial meniscotibial ligament, thereby destabilizing the joint and causing instability-driven OA. An important caveat for mouse models is that the mouse and human knee differ in terms of joint size, joint biomechanics, and histologic characteristics (layers, cellularity),^32,64 and joint differences could confound clinical translation.¹⁰ Table 1. Mouse models of osteoarthritis.