Immunomodulation by Mesenchymal Stem Cells in Veterinary Species

Mesenchymal stem cells (MSC) are adult-derived multipotent stem cells that have been derived from almost every tissue. They are classically defined as spindle-shaped, plastic-adherent cells capable of adipogenic, chondrogenic, and osteogenic differentiation. This capacity for trilineage differentiation has been the foundation for research into the use of MSC to regenerate damaged tissues. Recent studies have shown that MSC interact with cells of the immune system and modulate their function. Although many of the details underlying the mechanisms by which MSC modulate the immune system have been defined for human and rodent (mouse and rat) MSC, much less is known about MSC from other veterinary species. This knowledge gap is particularly important because the clinical use of MSC in veterinary medicine is increasing and far exceeds the use of MSC in human medicine. It is crucial to determine how MSC modulate the immune system for each animal species as well as for MSC derived from any given tissue source. A comparative approach provides a unique translational opportunity to bring novel cell-based therapies to the veterinary market as well as enhance the utility of animal models for human disorders. The current review covers what is currently known about MSC and their immunomodulatory functions in veterinary species, excluding laboratory rodents.Abbreviations: AT, adipose tissue; BM, Bone marrow; CB, umbilical cord blood; CT, umbilical cord tissue; DC, dendritic cell; IDO, indoleamine 2;3-dioxygenase; MSC, mesenchymal stem cells; PGE₂, prostaglandin E2; VEGF, vascular endothelial growth factorMesenchymal stem cells (MSC, alternatively known as mesenchymal stromal cells) were first reported in the literature in 1968.³⁹ MSC are thought to be of pericyte origin (cells that line the vasculature)^21,22 and typically are isolated from highly vascular tissues. In humans and mice, MSC have been isolated from fat, placental tissues (placenta, Wharton jelly, umbilical cord, umbilical cord blood), hair follicles, tendon, synovial membrane, periodontal ligament, and every major organ (brain, spleen, liver, kidney, lung, bone marrow, muscle, thymus, pancreas, skin).^23,121 For most current clinical applications, MSC are isolated from adipose tissue (AT), bone marrow (BM), umbilical cord blood (CB), and umbilical cord tissue (CT; 11,87,99 Clinical trials in human medicine focus on the use of MSC both for their antiinflammatory properties (graft-versus-host disease, irritable bowel syndrome) and their ability to aid in tissue and bone regeneration in combination with growth factors and bone scaffolds (clinicaltrials.gov).¹³¹ For tissue regeneration, the abilities of MSC to differentiate and to secrete mediators and interact with cells of the immune system likely contribute to tissue healing (Figure 1). The current review will not address the specific use of MSC for orthopedic applications and tissue regeneration, although the topic is covered widely in current literature for both human and veterinary medicine.^57,62,90

Table 1.

Tissues from which MSC have been isolated

Coordination of Leaf Photosynthesis,Transpiration, and Structural Traits in Rice and Wild Relatives (Genus Oryza)

The genus Oryza, which includes rice (Oryza sativa and Oryza glaberrima) and wild relatives, is a useful genus to study leaf properties in order to identify structural features that control CO₂ access to chloroplasts, photosynthesis, water use efficiency, and drought tolerance. Traits, 26 structural and 17 functional, associated with photosynthesis and transpiration were quantified on 24 accessions (representatives of 17 species and eight genomes). Hypotheses of associations within, and between, structure, photosynthesis, and transpiration were tested. Two main clusters of positively interrelated leaf traits were identified: in the first cluster were structural features, leaf thickness (Thick_leaf), mesophyll (M) cell surface area exposed to intercellular air space per unit of leaf surface area (S_mes), and M cell size; a second group included functional traits, net photosynthetic rate, transpiration rate, M conductance to CO₂ diffusion (g_m), stomatal conductance to gas diffusion (g_s), and the g_m/g_s ratio. While net photosynthetic rate was positively correlated with g_m, neither was significantly linked with any individual structural traits. The results suggest that changes in g_m depend on covariations of multiple leaf (S_mes) and M cell (including cell wall thickness) structural traits. There was an inverse relationship between Thick_leaf and transpiration rate and a significant positive association between Thick_leaf and leaf transpiration efficiency. Interestingly, high g_m together with high g_m/g_s and a low S_mes/g_m ratio (M resistance to CO₂ diffusion per unit of cell surface area exposed to intercellular air space) appear to be ideal for supporting leaf photosynthesis while preserving water; in addition, thick M cell walls may be beneficial for plant drought tolerance.Leaves have evolved in different environments into a multitude of sizes and shapes, showing great variation in morphology and anatomy (Evans et al., 2004). However, all leaf typologies share common functions associated with chloroplasts, namely to intercept sunlight, take up CO₂ and inorganic nitrogen, and perform photosynthesis as a primary process for growth and reproduction.Investigating relationships between leaf anatomy and photosynthetic features (CO₂ fixation, which involves physical and biochemical processes and loss of water by transpiration) could lead to the identification of structural features for enhancing crop productivity and improve our understanding of plant evolution and adaptation (Evans et al., 2004).Stomata, through which CO₂ and water vapor diffuse into and out of the leaf, are involved in the regulation and control of photosynthetic and transpiration responses (Jarvis and Morison, 1981; Farquhar and Sharkey, 1982). Besides stomata distribution patterns between the abaxial and adaxial lamina surfaces (Foster and Smith, 1986), stomatal density and size are leaf anatomical traits contributing to build the leaf stomatal conductance to gas diffusion (g_s). This is calculated as the reciprocal of the stomatal resistances to gas diffusion; stomatal control results in a lower concentration of CO₂ in the leaf mesophyll (M) intercellular air space (C_i) than in the atmosphere (C_a; Nobel, 2009).Leaf M architecture greatly contributes to the pattern of light attenuation profiles within the lamina (Terashima and Saeki, 1983; Woolley, 1983; Vogelmann et al., 1989; Evans, 1999; Terashima et al., 2011) and affects CO₂ diffusion from the intercellular air space (IAS) to the chloroplast stroma. Therefore, it influences photosynthetic activity (Flexas et al., 2007, 2008) and can have effects on leaf hydrology and transpiration (Sack et al., 2003; Brodribb et al., 2010; Ocheltree et al., 2012). In addition, M architecture sets boundaries for leaf photosynthetic responses to changing environmental conditions (Nobel et al., 1975).Fortunately, several methodologies are currently available (Flexas et al., 2008; Pons et al., 2009) to determine M conductance to CO₂ diffusion (g_m), expressed per unit of leaf surface area. It is calculated as the reciprocal of the cumulated partial resistances exerted by leaf structural traits and biochemical processes from the substomatal cavities to photosynthetic sites (Evans et al., 2009; Nobel, 2009). The resistance to CO₂ diffusion in the liquid phase is 4 orders of magnitude higher than in the gaseous phase (Nobel, 2009); therefore, the changes in CO₂ concentration in the leaf gas phase are small in comparison with the changes in the liquid phase (Niinemets, 1999; Aalto and Juurola, 2002; Nobel, 2009). In the liquid phase, the resistance to CO₂ transfer is built from contributions by the cell walls, the plasmalemma, cytoplasm, chloroplast membranes, and stroma (Tholen and Zhu, 2011; Tholen et al., 2012); in addition, it involves factors associated with the carboxylation reaction (Kiirats et al., 2002; Evans et al., 2009). Thus, the concentration of CO₂ in the chloroplasts (C_c) is lower than C_i and can limit photosynthesis.At steady state, the relationships between the leaf net photosynthetic rate (A), the concentrations of CO₂, and the stomatal conductance to CO₂ diffusion (g_{s_CO2}) and g_m are modeled based on Fick’s first law of diffusion (Nobel, 2009) as:(1)where C_a, C_i, and C_c are as defined above (Flexas et al., 2008).The magnitude of g_m has been found to correlate with certain leaf structural traits in some species, in particular with the M cell surface area exposed to IAS per (one side) unit of leaf surface area (S_mes) and its extent covered by chloroplasts (S_chl; Evans and Loreto, 2000; Slaton and Smith, 2002; Tholen et al., 2012). From a physical modeling perspective, increasing S_mes provides more pathways acting in parallel for CO₂ diffusion (to and from the chloroplasts) per unit of leaf surface area; thus, it tends to reduce the resistance to CO₂ movement into the M cells and to increase g_m (Evans et al., 2009; Nobel, 2009). A number of leaf structural traits affect S_mes, including leaf thickness, cell density, cell volume and shape, and the fraction of the M cell walls in contact with the IAS (Terashima et al., 2001, 2011), and the degree they are linked to S_mes can vary between species (Slaton and Smith, 2002; Terashima et al., 2006). In particular, the presence of lobes on M cells, which are prominent in some Oryza species, may contribute to g_m through increasing S_mes (Sage and Sage, 2009; Terashima et al., 2011; Tosens et al., 2012). The M cell wall can provide resistance in series for M CO₂ diffusion (Nobel, 2009); thicker cell walls may increase resistance to CO₂ movement into the M cells and decrease g_m (Terashima et al., 2006, 2011; Evans et al., 2009).Other leaf traits, such as M porosity (the fraction of M volume occupied by air spaces [Vol_IAS]), has been shown to have a positive correlation with g_m in some species (Peña-Rojas et al., 2005), but the association may be mediated by light availability (Slaton and Smith, 2002). Leaf thickness (Thick_leaf) tends to be negatively linked to g_m, and it may set an upper limit for the maximum g_m, according to Terashima et al. (2006), Flexas et al. (2008), and Niinemets et al. (2009).With respect to leaf structural traits and water relations, Thick_leaf may increase the apoplast path length (resistances in series; Nobel, 2009) in the extra-xylem M (Sack and Holbrook, 2006; Brodribb et al., 2007) for water to reach the evaporation sites, which could decrease the conductance of water through the M and lower the transpiration rate. Interestingly, while thicker M cell walls may reduce g_m, they can enable the development of higher water potential gradients between the soil and leaves, which can be decisive for plant survival and longevity under drought conditions (Steppe et al., 2011).The purpose of this study was to provide insight into how the diversity of leaf structure relates to photosynthesis and transpiration among representative cultivated species and wild relatives in the genus Oryza. This includes, in particular, identifying leaf structural features associated with the diffusion of CO₂ from the atmosphere to the chloroplasts, photosynthesis, transpiration efficiency (A/E), and drought tolerance. The genus consists of 10 genomic groups and is composed of approximately 24 species (the number depending on taxonomic preferences; Kellogg, 2009; Brar and Singh, 2011), including the cultivated species Oryza sativa and Oryza glaberrima. Oryza species are distributed around the world, and they exhibit a wide range of phenotypes, with annual versus perennial life cycles and sun- versus shade-adapted species (Vaughan, 1994; Vaughan et al., 2008; Brar and Singh, 2011; Jagadish et al., 2011). This diversity in the genus is an important resource, which is being studied to improve rice yield, especially under unfavorable environmental conditions. In particular, O. glaberrima, Oryza australiensis, and Oryza meridionalis are of interest as drought-tolerant species (Henry et al., 2010; Ndjiondjop et al., 2010; Scafaro et al., 2011, 2012), while Oryza coarctata is salt tolerant (Sengupta and Majumder, 2010). In this study, a total of 43 leaf functional and structural parameters were collected on 24 accessions corresponding to 17 species within eight genomes (Brar and Singh (2011). Life cycle is as follows: A = annual; B = biennial; P = poliennial. Habitat is as follows: S = shade; S-Sh = sun-shade.

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.

Natural Infection of Burkholderia pseudomallei in an Imported Pigtail Macaque (Macaca nemestrina) and Management of the Exposed Colony

Identification of the select agent Burkholderia pseudomallei in macaques imported into the United States is rare. A purpose-bred, 4.5-y-old pigtail macaque (Macaca nemestrina) imported from Southeast Asia was received from a commercial vendor at our facility in March 2012. After the initial acclimation period of 5 to 7 d, physical examination of the macaque revealed a subcutaneous abscess that surrounded the right stifle joint. The wound was treated and resolved over 3 mo. In August 2012, 2 mo after the stifle joint wound resolved, the macaque exhibited neurologic clinical signs. Postmortem microbiologic analysis revealed that the macaque was infected with B. pseudomallei. This case report describes the clinical evaluation of a B. pseudomallei-infected macaque, management and care of the potentially exposed colony of animals, and protocols established for the animal care staff that worked with the infected macaque and potentially exposed colony. This article also provides relevant information on addressing matters related to regulatory issues and risk management of potentially exposed animals and animal care staff.Abbreviations: CDC, Centers for Disease Control and Prevention; IHA, indirect hemagglutination assay; PEP, postexposure prophylacticBurkholderia pseudomallei, formerly known as Pseudomonas pseudomallei, is a gram-negative, aerobic, bipolar, motile, rod-shaped bacterium. B. pseudomallei infections (melioidosis) can be severe and even fatal in both humans and animals. This environmental saprophyte is endemic to Southeast Asia and northern Australia, but it has also been found in other tropical and subtropical areas of the world.^7,22,32,42 The bacterium is usually found in soil and water in endemic areas and is transmitted to humans and animals primarily through percutaneous inoculation, ingestion, or inhalation of a contaminated source.^{8, 22,28,32,42} Human-to-human, animal-to-animal, and animal-to-human spread are rare.^8,32 In December 2012, the National Select Agent Registry designated B. pseudomallei as a Tier 1 overlap select agent.³⁹ Organisms classified as Tier 1 agents present the highest risk of deliberate misuse, with the most significant potential for mass casualties or devastating effects to the economy, critical infrastructure, or public confidence. Select agents with this status have the potential to pose a severe threat to human and animal health or safety or the ability to be used as a biologic weapon.³⁹Melioidosis in humans can be challenging to diagnose and treat because the organism can remain latent for years and is resistant to many antibiotics.^12,37,41 B. pseudomallei can survive in phagocytic cells, a phenomenon that may be associated with latent infections.^19,38 The incubation period in naturally infected animals ranges from 1 d to many years, but symptoms typically appear 2 to 4 wk after exposure.^13,17,35,38 Disease generally presents in 1 of 2 forms: localized infection or septicemia.²² Multiple methods are used to diagnose melioidosis, including immunofluorescence, serology, and PCR analysis, but isolation of the bacteria from blood, urine, sputum, throat swabs, abscesses, skin, or tissue lesions remains the ‘gold standard.’^9,22,40,42 The prognosis varies based on presentation, time to diagnosis, initiation of appropriate antimicrobial treatment, and underlying comorbidities.^7,28,42 Currently, there is no licensed vaccine to prevent melioidosis.There are several published reports of naturally occurring melioidosis in a variety of nonhuman primates (NHP; 2,10,13,17,25,30,31,35 The first reported case of melioidosis in monkeys was recorded in 1932, and the first published case in a macaque species was in 1966.³⁰ In the United States, there have only been 7 documented cases of NHP with B. pseudomallei infection.^2,13,17 All of these cases occurred prior to the classification of B. pseudomallei as a select agent. Clinical signs in NHP range from subclinical or subacute illness to acute septicemia, localized infection, and chronic infection. NHP with melioidosis can be asymptomatic or exhibit clinical signs such as anorexia, wasting, purulent drainage, subcutaneous abscesses, and other soft tissue lesions. Lymphadenitis, lameness, osteomyelitis, paralysis and other CNS signs have also been reported.^{2,7,10,22,28,32} In comparison, human''s clinical signs range from abscesses, skin ulceration, fever, headache, joint pain, and muscle tenderness to abdominal pain, anorexia, respiratory distress, seizures, and septicemia.^7,9,21,22

Table 1.

Summary of reported cases of naturally occurring Burkholderia pseudomalleiinfections in nonhuman primates

Focus on Metabolism: Posttranslational Protein Modifications in Plant Metabolism

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

The Operation of Two Decarboxylases,Transamination, and Partitioning of C4 Metabolic Processes between Mesophyll and Bundle Sheath Cells Allows Light Capture To Be Balanced for the Maize C4 Pathway

The C₄ photosynthesis carbon-concentrating mechanism in maize (Zea mays) has two CO₂ delivery pathways to the bundle sheath (BS; via malate or aspartate), and rates of phosphoglyceric acid reduction, starch synthesis, and phosphoenolpyruvate regeneration also vary between BS and mesophyll (M) cells. The theoretical partitioning of ATP supply between M and BS cells was derived for these metabolic activities from simulated profiles of light penetration across a leaf, with a potential 3-fold difference in the fraction of ATP produced in the BS relative to M (from 0.29 to 0.96). A steady-state metabolic model was tested using varying light quality to differentially stimulate M or BS photosystems. CO₂ uptake, ATP production rate (J_ATP; derived with a low oxygen/chlorophyll fluorescence method), and carbon isotope discrimination were measured on plants under a low light intensity, which is considered to affect C₄ operating efficiency. The light quality treatments did not change the empirical ATP cost of gross CO₂ assimilation (J_ATP/GA). Using the metabolic model, measured J_ATP/GA was compared with the predicted ATP demand as metabolic functions were varied between M and BS. Transamination and the two decarboxylase systems (NADP-malic enzyme and phosphoenolpyruvate carboxykinase) were critical for matching ATP and reduced NADP demand in BS and M when light capture was varied under contrasting light qualities.Interest in the C₄ pathway has been increased by the potential for enhancing crop productivity and maintaining yield stability in the face of global warming and population pressure (Friso et al., 2010; Zhu et al., 2010; Covshoff and Hibberd, 2012). Maize (Zea mays), a C₄ plant of the NADP-malic enzyme (ME) subtype, is a leading grain production cereal (www.fao.org). C₄ photosynthesis is a shared activity between mesophyll (M; abbreviations are listed in BS) cells, coupled to allow the operation of a biochemical carbon-concentrating mechanism (CCM). The CCM effectively minimizes photorespiration by increasing the CO₂ concentration in the bundle sheath (C_BS), where Rubisco is exclusively expressed. Since BS and M are connected by plasmodesmata, some CO₂ retrodiffuses. The refixation of that escaping CO₂ by the CCM increases the activity of the CCM and the total ATP demand (ATP_BS + ATP_M) for gross CO₂ assimilation (GA; [ATP_BS + ATP_M]/GA), from a theoretical minimum of five ATPs (Furbank et al., 1990). Leakiness (Φ), the amount of CO₂ retrodiffusing relative to phosphoenolpyruvate (PEP) carboxylation rate, is therefore a proxy for the coordination between the CCM and assimilatory activity (Henderson et al., 1992; Tazoe et al., 2008; Kromdijk et al., 2010; Ubierna et al., 2011; Bellasio and Griffiths, 2013).

Table I.

Variables and acronyms described in the text

Focus Issue on Roots: The Optimal Lateral Root Branching Density for Maize Depends on Nitrogen and Phosphorus Availability

Observed phenotypic variation in the lateral root branching density (LRBD) in maize (Zea mays) is large (1–41 cm⁻¹ major axis [i.e. brace, crown, seminal, and primary roots]), suggesting that LRBD has varying utility and tradeoffs in specific environments. Using the functional-structural plant model SimRoot, we simulated the three-dimensional development of maize root architectures with varying LRBD and quantified nitrate and phosphorus uptake, root competition, and whole-plant carbon balances in soils varying in the availability of these nutrients. Sparsely spaced (less than 7 branches cm⁻¹), long laterals were optimal for nitrate acquisition, while densely spaced (more than 9 branches cm⁻¹), short laterals were optimal for phosphorus acquisition. The nitrate results are mostly explained by the strong competition between lateral roots for nitrate, which causes increasing LRBD to decrease the uptake per unit root length, while the carbon budgets of the plant do not permit greater total root length (i.e. individual roots in the high-LRBD plants stay shorter). Competition and carbon limitations for growth play less of a role for phosphorus uptake, and consequently increasing LRBD results in greater root length and uptake. We conclude that the optimal LRBD depends on the relative availability of nitrate (a mobile soil resource) and phosphorus (an immobile soil resource) and is greater in environments with greater carbon fixation. The median LRBD reported in several field screens was 6 branches cm⁻¹, suggesting that most genotypes have an LRBD that balances the acquisition of both nutrients. LRBD merits additional investigation as a potential breeding target for greater nutrient acquisition.At least four major classes of plant roots can be distinguished based on the organ from which they originate: namely the seed, the shoot, the hypocotyl/mesocotyl, and other roots (Zobel and Waisel, 2010). The last class is lateral roots, which form in most plants the majority of the root length, but not necessarily of the root weight, as lateral roots have smaller diameter. Lateral roots start with the formation of lateral root primordia, closely behind the root tip of the parent root. These primordia undergo nine distinguishable steps, of which the last step is the emergence from the cortex of the parent root just behind the zone of elongation, usually only a few days after the first cell divisions that lead to their formation (Malamy and Benfey, 1997). However, not all primordia develop into lateral roots; some stay dormant (Dubrovsky et al., 2006), although dormancy of primordia may not occur in maize (Zea mays; Jordan et al., 1993; Ploshchinskaia et al., 2002). The final number of lateral roots is thereby dependent on the rate of primordia formation as well as the percentage of primordia that develop into lateral roots. This process of primordia formation and lateral root emergence is being studied intensively, including the genes that are activated during the different steps and the hormones regulating the process (López-Bucio et al., 2003; Dubrovsky et al., 2006; Osmont et al., 2007; Péret et al., 2009; Lavenus et al., 2013). Significant genotypic variation in the density of lateral roots has been observed, ranging from no lateral roots to 41 roots cm⁻¹ in maize (Trachsel et al., 2010; Lynch, 2013). This suggests that clear tradeoffs exist for the development of lateral roots and that these genotypes have preprogrammed growth patterns that are adaptive to specific environments. While some of the variation for lateral root branching density (LRBD) that has been observed across environments, for example by Trachsel et al. (2010), is constitutive, many genotypes have strong plasticity responses of LRBD to variations in soil fertility (Zhu et al., 2005a; Osmont et al., 2007). Both the nutrient and carbon status of the plant and the local nutrient environment of the (parent) root tip influence LRBD. Many studies have documented these plasticity responses, and others have tried to unravel parts of the sensing and signaling pathways that regulate LRBD. The utility of root proliferation into a nutrient patch has been studied and debated (Robinson et al., 1999; Hodge, 2004), but much less so the utility of having fewer or more branches across the whole root system. Our understanding of the adaptive significance of variation in LRBD among genotypes is thereby limited, with many studies not accounting for relevant tradeoffs. In this study, we integrate several functional aspects of LRBD with respect to nutrient acquisition, root competition, and internal resource costs and quantify these functional aspects using the functional structural plant model SimRoot. SimRoot simulates plant growth with explicit representation of root architecture in three dimensions (Fig. 1; Supplemental Movie S1). The model focuses on the resource acquisition by the root system and carbon fixation by the shoot while estimating the resource utilization and requirements by all the different organs.

Table I.

Minimum, maximum, and median LRBD in different populations phenotyped by various researchers at several locations in the worldLocations are as follows: D, Juelich, Germany; PA, State College, PA; and SA, Alma, South Africa. Some of the experiments included nutrient treatments: LN, low nitrogen availability; and LP, low phosphorus availability. Data were collected by counting the number of roots along a nodal root segment. Data were supplied by the person indicated under source: H.S., H. Schneider; L.Y., L. York; A.Z., A. Zhan; and J.P., J.A. Postma. WiDiv, Wisconsin Diversity panel; IBM, intermated B73 × Mo17; NAM, nested association mapping.

Rethinking Guard Cell Metabolism

Stomata control gaseous fluxes between the internal leaf air spaces and the external atmosphere and, therefore, play a pivotal role in regulating CO₂ uptake for photosynthesis as well as water loss through transpiration. Guard cells, which flank the stomata, undergo adjustments in volume, resulting in changes in pore aperture. Stomatal opening is mediated by the complex regulation of ion transport and solute biosynthesis. Ion transport is exceptionally well understood, whereas our knowledge of guard cell metabolism remains limited, despite several decades of research. In this review, we evaluate the current literature on metabolism in guard cells, particularly the roles of starch, sucrose, and malate. We explore the possible origins of sucrose, including guard cell photosynthesis, and discuss new evidence that points to multiple processes and plasticity in guard cell metabolism that enable these cells to function effectively to maintain optimal stomatal aperture. We also discuss the new tools, techniques, and approaches available for further exploring and potentially manipulating guard cell metabolism to improve plant water use and productivity.Stomata are microscopic, adjustable pores on the leaf surface. The evolution of stomata more than 400 million years ago (Edwards et al., 1986, 1992, 1998) helped facilitate the adaptation of plants to a terrestrial environment, where water is typically a limiting resource. Each stoma is composed of two kidney- or dumbbell-shaped guard cells, whose volume changes to adjust pore aperture, allowing plants to simultaneously regulate CO₂ uptake and water loss. This facilitation of gas exchange by stomatal opening is one of the most essential processes in plant photosynthesis and transpiration, affecting plant water use efficiency and agricultural crop yields (Lawson and Blatt, 2014).Plant physiologists have a long history of investigating the behavior of these fascinating structures, reaching back more than a century to the pioneering work of Sir Francis Darwin (Darwin, 1916) and the American botanist Francis Ernest Lloyd (Lloyd, 1908). Major contributions to stomatal research arose from inventing and improving equipment and methods for quantitatively measuring the effects of environmental factors on stomatal pore aperture. After Darwin’s work, it became clear that the stomatal aperture actively responds to changes in the environment and regulates leaf transpiration rates (Meidner, 1987). Over the past century, much has been learned about their structure, development, and physiology.Despite the anatomical simplicity of the stomatal valve, the surrounding guard cells are highly specialized. Guard cells are morphologically distinct from general epidermal cells and possess complex signal transduction networks, elevated membrane ion transport capacity, and modified metabolic pathways. These features allow rapid modulations in guard cell turgor in response to endogenous and environmental signals, promoting the opening and closure of the stomatal pore in time scales of seconds to hours (Assmann and Wang, 2001). A variety of osmotically active solutes contribute to the buildup of stomatal turgor. Potassium (K⁺) and chloride (Cl⁻) act as the main inorganic ions, and malate²⁻ and sucrose (Suc) function as the main organic solutes. Whereas K⁺ and Cl⁻ are taken up from the apoplast, Suc and malate²⁻ can be imported or synthesized internally using carbon skeletons deriving from starch degradation and/or CO₂ fixation in the guard cell chloroplast (Roelfsema and Hedrich, 2005; Vavasseur and Raghavendra, 2005; Lawson, 2009; Kollist et al., 2014). The accumulation of these osmotica lowers the water potential, promoting the inflow of water, the swelling of guard cells, and the opening of the stomatal pore. Most of the ions taken up, or synthesized by guard cells, are sequestered into the vacuole (Barbier-Brygoo et al., 2011). As a result, the guard cell vacuoles undergo dynamic changes in volume and structure, which are crucial for achieving the full amplitude of stomatal movements (Gao et al., 2005; Tanaka et al., 2007; Andrés et al., 2014). During stomatal closure, guard cells reduce their volume through the release of ions into the cell wall and the consequent efflux of water.The transport of osmolytes across the plasma and tonoplast guard cell membranes is energized by H⁺-ATPase activity, which generates a proton motive force by translocating H⁺ ions against their concentration gradient (Blatt, 1987a, 1987b; Thiel et al., 1992; Roelfsema and Hedrich, 2005; Gaxiola et al., 2007). After the pioneering work of Fischer demonstrated the importance of K⁺ uptake in stomatal opening (Fischer, 1968; Fischer and Hsiao, 1968), K⁺ transport became of central interest and has long been considered the essence of stomatal movement regulation. The development of the voltage clamp technique, along with the relative easy acquisition of knockout mutants and transgenics in the model plant Arabidopsis (Arabidopsis thaliana), helped to uncover the precise mechanism and function of K⁺ fluxes in guard cells. It is well established that changes in membrane potential in response to several stimuli (e.g. light/darkness, CO₂, and abscisic acid [ABA]) alter the direction of K⁺ transport (Thiel et al., 1992; Blatt, 2000; Roelfsema et al., 2001, 2002, 2004). During stomatal opening, the activation of the proton pump generates a sufficiently negative electric potential to cause the uptake of K⁺ through the inward-rectifying K⁺ channels (K⁺_in; Fig. 1). During stomatal closure, K⁺ outflow from outward-rectifying K⁺ channels (K⁺_out) results from membrane depolarization (Fig. 2; Blatt, 1988; Schroeder, 1988; Anderson et al., 1992; Sentenac et al., 1992). Besides being gated by opposing changes in voltage, the activation of (K⁺_out) channels is dependent on the extracellular K⁺ concentration, while that of K⁺_in is not (Blatt, 1988, 1992; Roelfsema and Prins, 1997; Dreyer and Blatt, 2009). There is also strong evidence for H⁺-coupled K⁺ symport in guard cells, which could account for up to 50% of total K⁺ uptake during stomatal opening (Blatt and Clint, 1989; Clint and Blatt, 1989; Hills et al., 2012). At least for K⁺_in, the loss of a single-channel gene in Arabidopsis has little or no impact on stomatal movement (Szyroki et al., 2001), showing the redundancy among the different K⁺_in isoforms and of K⁺ transport in general.Open in a separate windowFigure 1.Integration of guard cell carbohydrate metabolism with membrane ion transport during stomatal opening. Sugars in guard cells can be imported from the apoplast, derive from starch breakdown, or be synthesized in the Calvin cycle. These sugars then can be stored as osmotically active solutes in the vacuole or metabolized in the cytosol to yield energy, reducing equivalents, and phosphoenolpyruvate (PEP). PEP can be further metabolized to pyruvate in the mitochondrial tricarboxylic acid (CAC) cycle or used as carbon skeletons for the biosynthesis of malate via PEP carboxylase (PEPC) and NAD-dependent malate dehydrogenase (NAD-MDH). Malate (which also can be imported from the apoplast) and the inorganic ions K⁺ and Cl⁻ accumulate in the vacuole, lowering the guard cell osmotic potential, thereby promoting stomatal opening. ABCB14, ATP-binding cassette transporter B14; AcetylCoA, acetyl-CoA; ALMT, aluminum-activated malate transporter; ATP-PFK, ATP-dependent phosphofructokinase; AttDT, dicarboxylate transporter; cINV, cytosolic invertase; cwINV, cell wall invertase; Fru6P, Fru-6-P; Fru1,6P₂, fructose 1,6-bisphosphate; Gl6P, Glc-6-P; G3P, glyceraldehyde 3-phosphate; iPGAM, phosphoglycerate mutase isoforms; NRGA1, negative regulator of guard cell ABA signaling1; OAA, oxaloacetate; 2-PGA, 2-phosphoglycerate; 3-PGA, 3-phosphoglycerate; PPi-PFK, PPi-dependent Fru-6-P phosphotransferase; STP, monosaccharide/H⁺ cotransporter; SUC, Suc/H⁺ cotransporter; SuSy, Suc synthase; TPT, triose phosphate/phosphate translocator. Compartments are not to scale. The dotted line indicates multiple metabolic steps.Open in a separate windowFigure 2.Proposed pathways of osmolyte dissipation during stomatal closure. While the removal of Cl⁻ and K⁺ is well described in the literature, the fate of Suc and malate during stomatal closure is unclear. Suc can be cleaved by cytosolic invertase (cINV), and the resulting hexoses can be imported into the chloroplast in the form of Glc-6-P (Glc6P). Glc6P is used subsequently for starch biosynthesis. Malate can be removed from the cell via decarboxylation to pyruvate by malic enzyme (ME) and the subsequent complete oxidation in the mitochondrial tricarboxylic acid (CAC) cycle. Alternatively, malate can be converted to PEP via NAD⁺-dependent malate dehydrogenase (NAD-MDH) and PEP carboxykinase (PEPCK). Gluconeogenic conversion of PEP to Glc6P establishes a possible link between malate removal and starch synthesis. Compartments are not to scale. PEP, Phosphoenolpyruvate; OAA; oxaloacetate; STP, monosaccharide/H⁺ cotransporter; SUC, Suc/H⁺ cotransporter; SuSy, Suc synthase; cINV, cytosolic invertase; NRGA1, negative regulator of guard cell ABA signaling1; ALMT, aluminum-activated malate transporter; GPT, Glc-6-P/Pi translocator; cwINV, cell wall invertase; HK, hexokinase; QUAC1, quickly activating anion channel1.Despite the undisputed importance of K⁺ uptake in stomatal opening, the accumulation of K⁺ ions alone cannot account for the increase in osmotic pressure necessary to explain stomatal aperture. Studies from the 1980s by MacRobbie and Fischer demonstrated that Vicia faba guard cells take up approximately 2 pmol of K⁺ during stomatal opening. Assuming that K⁺ uptake is balanced by the accumulation of similar amounts of counter ions (Cl⁻ and/or malate²⁻), the expected increase in stomatal turgor to approximately 3 MPa is less than the 4.5 MPa expected for fully open stomata (Fischer, 1972; MacRobbie and Lettau, 1980a, 1980b; Chen et al., 2012). The realization that other solutes must accumulate in addition to K⁺ salts was one of the major paradigm shifts in stomatal physiology research in the last decades, equal to the discovery of ion channels. Suc was put forward as the most likely candidate for the additional osmoticum to support stomatal opening (MacRobbie, 1987; Tallman and Zeiger, 1988; Talbott and Zeiger, 1993, 1998). Nonetheless, this research area subsequently failed to attract notice commensurate with its importance.In the last few years, the metabolism of starch, sugars and, organic acids in guard cells has seen a rebirth, making this the perfect time to review the developments in this field. In this review, we focus on photosynthetic carbon assimilation and respiratory metabolism in guard cells and provide a historical overview of the subject that highlights the most up-to-date and novel discoveries in guard cell research. We describe the various metabolic pathways separately, but as metabolism is an integrated network, we also discuss their reciprocal and beneficial interactions. Finally, we highlight their connection with the metabolism in the subjacent mesophyll cells and how they integrate with guard cell signal transduction networks and membrane ion transport to regulate stomatal movements. The enzymes and transporters discussed in this review are listed in Arabidopsis Genome Initiative CodeGeneProteinFunctionMalate transport AT1G28010ABCB14ATP-binding cassette transporter B14Import of apoplastic malate AT5G47560tDTDicarboxylate transporterTransport of carboxylates into the vacuole AT3G18440ALMT9Aluminum-activated malate transporter9Transport of Cl⁻/malate²⁻ into the vacuole AT2G17470ALMT6Aluminum-activated malate transporter6Transport of malate²⁻ into the vacuole AT4G17970ALMT12/QUAC1Aluminum-activated malate transporter12Export of cytosolic Cl⁻/malate²⁻ to the apoplastMalate metabolism –PEPCPhosphoenolpyruvate carboxylaseβ-Carboxylation of PEP to OAA –NAD-MDHNAD⁺-dependent malate dehydrogenaseReduction of OAA to malate –MEMalic enzymeOxidative decarboxylation of malate to pyruvate AT4G37870PEPCK1PEP carboxykinase1Conversion of OAA to PEP –PPDKPyruvate, orthophosphate dikinaseConversion of pyruvate to PEPOther carboxylates –TPTTriose phosphate/phosphate translocatorExport of triose phosphate from the chloroplast to the cytosol AT4G05590NRGA1Negative regulator of guard cell ABA signaling1Putative mitochondrial pyruvate carrier –SDH2Succinate dehydrogenase2Oxidation of succinate to fumarate AT2G47510FUM1Fumarase1Hydration of fumarate to malate –iPGAMPhosphoglycerate mutaseInterconversion of 3-PGA to 2-PGA –PPi-PFKPPi-dependent Fru-6-P phosphotransferasePhosphorylation of Fru-6-P to Fru-1,6-bisphosphate –ATP-PFKATP-dependent phosphofructokinasePhosphorylation of Fru-6-P to Fru-1,6-bisphosphateCalvin cycle –RubiscoRubiscoCarboxylation of ribulose 1,5-bisphosphate AT3G55800SBPaseSedoheptulose-bisphosphataseDephosphorylation of sedoheptulose-1,7-bisphosphate to sedoheptulose-7-phosphateSugar metabolism AT4G29130HK1Hexokinase1Phosphorylation of Glc to Glc-6-P AT4G02280SuSySuc synthase3Interconversion of Suc to Fru and UDP-Glc –cINVCytosolic invertaseHydrolysis of Suc to Fru and Glc –cwINVCell wall invertaseHydrolysis of Suc to Fru and GlcSugar transport AT1G11260STP1Monosaccharide/H⁺ cotransporter1Import of apoplastic hexose sugars AT3G19930STP4Monosaccharide/H⁺ cotransporter4Import of apoplastic hexose sugars AT1G71880SUC1Suc/H⁺ cotransporter1Import of apoplastic Suc AT2G02860SUC3Suc/H⁺ cotransporter3Import of apoplastic SucStarch degradation AT3G23920BAM1β-Amylase1Hydrolysis of α-1,4 external glucoside linkages in starch AT1G69830AMY3α-Amylase3Hydrolysis of α-1,4 internal glucoside linkages in starchStarch synthesis –GPTGlc-6-P/Pi translocatorUptake of cytosolic Glc-6-P into the chloroplast AT4G24620PGIPhosphoglucose isomeraseConversion of Fru-6-P to Glc-6-P AT5G51820PGM1Phosphoglucomutase1Conversion of Glc-6-P to Glc-1-P AT5G48300APS1ADPGlc pyrophosphorylase small subunitConversion of Glc-1-P to ADPGlc, catalytic subunit –APLADPGlc pyrophosphorylase large subunitConversion of Glc-1-P to ADPGlc, regulatory subunitVarious AT3G45780PHOT1Phototropin1Blue light photoreceptor AT5G58140PHOT2Phototropin2Blue light photoreceptor AT4G14480BLUS1Blue light signaling1Protein kinase, regulator of blue light-induced stomatal opening –PP1Protein phosphatase1Regulator of blue light-induced stomatal opening AT3G01500CA1Carbonic anhydrase1Interconversion of CO₂ and water into H₂CO₃ AT1G70410CA4Carbonic anhydrase4Interconversion of CO₂ and water into H₂CO₃ AT1G62400HT1High leaf temperature1Protein kinase, regulator of CO₂-induced stomatal closureOpen in a separate window     

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.     

Importin-α-Mediated Nucleolar Localization of Potato Mop-Top Virus TRIPLE GENE BLOCK1 (TGB1) Protein Facilitates Virus Systemic Movement,Whereas TGB1 Self-Interaction Is Required for Cell-to-Cell Movement in Nicotiana benthamiana

Recently, it has become evident that nucleolar passage of movement proteins occurs commonly in a number of plant RNA viruses that replicate in the cytoplasm. Systemic movement of Potato mop-top virus (PMTV) involves two viral transport forms represented by a complex of viral RNA and TRIPLE GENE BLOCK1 (TGB1) movement protein and by polar virions that contain the minor coat protein and TGB1 attached to one extremity. The integrity of polar virions ensures the efficient movement of RNA-CP, which encodes the virus coat protein. Here, we report the involvement of nuclear transport receptors belonging to the importin-α family in nucleolar accumulation of the PMTV TGB1 protein and, subsequently, in the systemic movement of the virus. Virus-induced gene silencing of two importin-α paralogs in Nicotiana benthamiana resulted in significant reduction of TGB1 accumulation in the nucleus, decreasing the accumulation of the virus progeny in upper leaves and the loss of systemic movement of RNA-CP. PMTV TGB1 interacted with importin-α in N. benthamiana, which was detected by bimolecular fluorescence complementation in the nucleoplasm and nucleolus. The interaction was mediated by two nucleolar localization signals identified by bioinformatics and mutagenesis in the TGB1 amino-terminal domain. Our results showed that while TGB1 self-interaction is needed for cell-to-cell movement, importin-α-mediated nucleolar targeting of TGB1 is an essential step in establishing the efficient systemic infection of the entire plant. These results enabled the identification of two separate domains in TGB1: an internal domain required for TGB1 self-interaction and cell-to-cell movement and the amino-terminal domain required for importin-α interaction in plants, nucleolar targeting, and long-distance movement.Pomoviruses are causal agents of important diseases affecting potato (Solanum tuberosum), sugar beet (Beta vulgaris), and bean (Phaseolus vulgaris). Potato mop-top virus (PMTV), the type member of the genus Pomovirus, causes an economically important disease of potato called spraing, inducing brown lines and arcs internally and on the surface of tubers. PMTV is transmitted by the root- and tuber-infecting plasmodiophorid Spongospora subterranea (Jones and Harrison, 1969; Arif et al., 1995).The pomovirus genome is divided into three single-stranded RNA (ssRNA) segments of positive polarity. RNA-Rep encodes the putative RNA-dependent RNA polymerase, the replicase of the virus (Savenkov et al., 1999). RNA-CP encodes a coat protein (CP) and another protein called CP-RT or minor CP, which is produced by translational read-through of the CP stop codon (Sandgren et al., 2001). Whereas CP is the major structural protein of the virions, CP-RT is incorporated in one of the termini of the virus particles and a domain within the read-through region of the protein is needed for transmission of the virus by its vector (Reavy et al., 1998). Moreover, CP-RT, but not CP, interacts with the major movement protein TRIPLE GENE BLOCK1 (TGB1; Torrance et al., 2009), which is encoded by RNA-TGB. Besides encoding a triple gene block of movement proteins, TGB1, TGB2, and TGB3 (Zamyatnin et al., 2004), RNA-TGB also encodes a viral suppressor of RNA silencing, the 8K protein (Lukhovitskaya et al., 2013b).To establish a successful infection in the entire plant, viruses must be able to replicate and to move their genomic components between cells, tissues, and organs. Recently, it has become evident that PMTV utilizes a sophisticated mode of cell-to-cell and long-distance movement that involves two virus transport forms, one represented by the viral nucleoprotein complexes (vRNPs) consisting of virus RNA and the TGB1 protein and another represented by the polar virions containing CP-RT and TGB1 proteins attached to one extremity of virus particles (Torrance et al., 2009; for review, see Solovyev and Savenkov, 2014). Proteins implicated in PMTV cell-to-cell movement include TGB1, TGB2, and TGB3 (Zamyatnin et al., 2004; Haupt et al., 2005a). Indirect evidence suggests that CP-RT is required for the efficient systemic movement of intact virions through its interaction with TGB1 (Torrance et al., 2009).Early in infection, the vRNP is transported on the endoplasmic reticulum actomyosin network and targeted to plasmodesmata by TGB2 and TGB3. Later in infection, fluorescently labeled TGB1 is seen in the nucleus and accumulates in the nucleolus. Nucleolar TGB1 association has been shown to be necessary for long-distance movement (Wright et al., 2010).Two structurally distinct subdomains have been identified in the N terminus of TGB1 proteins of hordeiviruses and pomoviruses (Makarov et al., 2009), an N-terminal domain (NTD) comprising approximately 125 amino acids in PMTV (ssRNA in noncooperative and cooperative manners, respectively. The C-terminal half of TGB1 contains a nucleoside triphosphatase/helicase domain that displays cooperative RNA binding. Previously, Wright et al. (2010) reported that TGB1 expressed from a 35S promoter localizes in the cytoplasm and accumulates in the nucleus and nucleolus with occasional labeling of microtubules (MTs). The MT labeling was apparent behind the leading edge of infection when yellow fluorescent protein (YFP)-TGB1 was expressed from an infectious clone. Deletion of 84 amino acids from the N terminus of TGB1 (representing most of the NTD) resulted in the absence of MTs, and nucleolar labeling and fusion of these 84 N-terminal amino acids to GFP resulted in nucleolar enrichment of GFP but no labeling of MTs. Deletion of the 5′ proximal part of the TGB1 open reading frame (ORF), encoding this N-terminal 84 amino acids, in the virus clone abolished systemic but not cell-to-cell movement. However, such deletion had no effect on TGB1 interactions with the CP-RT or self-interaction (Wright et al., 2010).

Table I.

Structural features of the PMTV TGB1 proteinPositively charged amino acids are set in boldface type and underscored. NoD, Nucleolar localization sequence detector; NS, not shown.