13.14: Smooth Muscle - Biology

13.14: Smooth Muscle - Biology

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Learning Objectives

  • Describe a dense body
  • Explain how smooth muscle works with internal organs and passageways through the body
  • Explain how smooth muscles differ from skeletal and cardiac muscles
  • Explain the difference between single-unit and multi-unit smooth muscle

Smooth muscle (so-named because the cells do not have striations) is present in the walls of hollow organs like the urinary bladder, uterus, stomach, intestines, and in the walls of passageways, such as the arteries and veins of the circulatory system, and the tracts of the respiratory, urinary, and reproductive systems (Figure 1). Smooth muscle is also present in the eyes, where it functions to change the size of the iris and alter the shape of the lens; and in the skin where it causes hair to stand erect in response to cold temperature or fear.

View the University of Michigan WebScope to explore the tissue sample in greater detail.

Smooth muscle fibers are spindle-shaped (wide in the middle and tapered at both ends, somewhat like a football) and have a single nucleus; they range from about 30 to 200 μm (thousands of times shorter than skeletal muscle fibers), and they produce their own connective tissue, endomysium. Although they do not have striations and sarcomeres, smooth muscle fibers do have actin and myosin contractile proteins, and thick and thin filaments. These thin filaments are anchored by dense bodies. A dense body is analogous to the Z-discs of skeletal and cardiac muscle fibers and is fastened to the sarcolemma. Calcium ions are supplied by the SR in the fibers and by sequestration from the extracellular fluid through membrane indentations called calveoli.

Because smooth muscle cells do not contain troponin, cross-bridge formation is not regulated by the troponin-tropomyosin complex but instead by the regulatory protein calmodulin. In a smooth muscle fiber, external Ca++ ions passing through opened calcium channels in the sarcolemma, and additional Ca++ released from SR, bind to calmodulin. The Ca++-calmodulin complex then activates an enzyme called myosin (light chain) kinase, which, in turn, activates the myosin heads by phosphorylating them (converting ATP to ADP and Pi, with the Pi attaching to the head). The heads can then attach to actin-binding sites and pull on the thin filaments. The thin filaments also are anchored to the dense bodies; the structures invested in the inner membrane of the sarcolemma (at adherens junctions) that also have cord-like intermediate filaments attached to them.

When the thin filaments slide past the thick filaments, they pull on the dense bodies, structures tethered to the sarcolemma, which then pull on the intermediate filaments networks throughout the sarcoplasm. This arrangement causes the entire muscle fiber to contract in a manner whereby the ends are pulled toward the center, causing the midsection to bulge in a corkscrew motion (Figure 2).

Although smooth muscle contraction relies on the presence of Ca++ ions, smooth muscle fibers have a much smaller diameter than skeletal muscle cells. T-tubules are not required to reach the interior of the cell and therefore not necessary to transmit an action potential deep into the fiber. Smooth muscle fibers have a limited calcium-storing SR but have calcium channels in the sarcolemma (similar to cardiac muscle fibers) that open during the action potential along the sarcolemma. The influx of extracellular Ca++ ions, which diffuse into the sarcoplasm to reach the calmodulin, accounts for most of the Ca++ that triggers contraction of a smooth muscle cell.

Muscle contraction continues until ATP-dependent calcium pumps actively transport Ca++ ions back into the SR and out of the cell. However, a low concentration of calcium remains in the sarcoplasm to maintain muscle tone. This remaining calcium keeps the muscle slightly contracted, which is important in certain tracts and around blood vessels.

Because most smooth muscles must function for long periods without rest, their power output is relatively low, but contractions can continue without using large amounts of energy. Some smooth muscle can also maintain contractions even as Ca++ is removed and myosin kinase is inactivated/dephosphorylated. This can happen as a subset of cross-bridges between myosin heads and actin, called latch-bridges, keep the thick and thin filaments linked together for a prolonged period, and without the need for ATP. This allows for the maintaining of muscle “tone” in smooth muscle that lines arterioles and other visceral organs with very little energy expenditure.

Smooth muscle is not under voluntary control; thus, it is called involuntary muscle. The triggers for smooth muscle contraction include hormones, neural stimulation by the ANS, and local factors. In certain locations, such as the walls of visceral organs, stretching the muscle can trigger its contraction (the stretch-relaxation response).

Axons of neurons in the ANS do not form the highly organized NMJs with smooth muscle, as seen between motor neurons and skeletal muscle fibers. Instead, there is a series of neurotransmitter-filled bulges called varicosities as an axon courses through smooth muscle, loosely forming motor units (Figure 3). A varicosity releases neurotransmitters into the synaptic cleft. Also, visceral muscle in the walls of the hollow organs (except the heart) contains pacesetter cells. A pacesetter cell can spontaneously trigger action potentials and contractions in the muscle.

Smooth muscle is organized in two ways: as single-unit smooth muscle, which is much more common; and as multiunit smooth muscle. The two types have different locations in the body and have different characteristics.

Single-unit muscle has its muscle fibers joined by gap junctions so that the muscle contracts as a single unit. This type of smooth muscle is found in the walls of all visceral organs except the heart (which has cardiac muscle in its walls), and so it is commonly called visceral muscle. Because the muscle fibers are not constrained by the organization and stretchability limits of sarcomeres, visceral smooth muscle has a stress-relaxation response. This means that as the muscle of a hollow organ is stretched when it fills, the mechanical stress of the stretching will trigger contraction, but this is immediately followed by relaxation so that the organ does not empty its contents prematurely. This is important for hollow organs, such as the stomach or urinary bladder, which continuously expand as they fill. The smooth muscle around these organs also can maintain a muscle tone when the organ empties and shrinks, a feature that prevents “flabbiness” in the empty organ. In general, visceral smooth muscle produces slow, steady contractions that allow substances, such as food in the digestive tract, to move through the body.

Multiunit smooth muscle cells rarely possess gap junctions, and thus are not electrically coupled. As a result, contraction does not spread from one cell to the next, but is instead confined to the cell that was originally stimulated. Stimuli for multiunit smooth muscles come from autonomic nerves or hormones but not from stretching. This type of tissue is found around large blood vessels, in the respiratory airways, and in the eyes.

Hyperplasia in Smooth Muscle

Similar to skeletal and cardiac muscle cells, smooth muscle can undergo hypertrophy to increase in size. Unlike other muscle, smooth muscle can also divide to produce more cells, a process called hyperplasia. This can most evidently be observed in the uterus at puberty, which responds to increased estrogen levels by producing more uterine smooth muscle fibers, and greatly increases the size of the myometrium.

Vascular smooth muscle cells in atherosclerosis

Vascular smooth muscle cells (VSMCs) are a major cell type present at all stages of an atherosclerotic plaque. According to the 'response to injury' and 'vulnerable plaque' hypotheses, contractile VSMCs recruited from the media undergo phenotypic conversion to proliferative synthetic cells that generate extracellular matrix to form the fibrous cap and hence stabilize plaques. However, lineage-tracing studies have highlighted flaws in the interpretation of former studies, revealing that these studies had underestimated both the content and functions of VSMCs in plaques and have thus challenged our view on the role of VSMCs in atherosclerosis. VSMCs are more plastic than previously recognized and can adopt alternative phenotypes, including phenotypes resembling foam cells, macrophages, mesenchymal stem cells and osteochondrogenic cells, which could contribute both positively and negatively to disease progression. In this Review, we present the evidence for VSMC plasticity and summarize the roles of VSMCs and VSMC-derived cells in atherosclerotic plaque development and progression. Correct attribution and spatiotemporal resolution of clinically beneficial and detrimental processes will underpin the success of any therapeutic intervention aimed at VSMCs and their derivatives.

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Beggs Laboratory | Muscle Biology 101

This section contains information about different muscle types, with emphasis on skeletal muscle. We will also go over the structure and function of muscle tissue. Finally, as muscle composition is determined by our genes, a brief introduction to certain genetics concepts will be presented.

Top to bottom left to right 1) Sanger sequencing showing the individual nucleotide base calls St 2) The 3 types of muscle cells 3) Muscle Structure 4) What cardiac muscle cells look like under magnification.

Muscle tissue is specialized for contraction and movement and it is found in two main forms: smooth and striated. Smooth muscle is present in those body systems that are under involuntary control. The digestive tract and the respiratory passages, among others, are made primarily of smooth muscle. Striated muscle is present in the heart and in the muscles that control our movements and breathing. Striated muscle owes its name to the interesting array of bands that becomes visible at the microscopic level. The origin and importance of these bands will be discussed further into this section. Striated muscle can be divided into two sub-types: cardiac muscle and skeletal muscle. Cardiac muscle , as the word implies, makes up the walls of the heart. Skeletal muscle is connected with the nerves, and because movement of these muscles can be consciously controlled, it is known as voluntary muscle. Most skeletal muscle is attached to our skeleton (hence its name) by a special form of tissue called tendons. In the Beggs Laboratory , we focus on the study of conditions that are caused by primary defects of skeletal muscle structure and function. These disorders are also known as myopathies .

Like any type of muscle, skeletal muscle is made of specialized cells called myofibers . Myofibers contain bundles of even smaller, thread-like structures called myofibrils . The myofibrils make up the machinery that allows movement and the use of force in daily activities. They contain many different specialized proteins that assemble together in a highly organized manner to form "thick filaments" and "thin filaments". The interaction between the proteins that form the thick and the thin filaments is what generates muscle contraction.

In summary, skeletal muscle is made of fibers, fibers are made of myofibrils, and myofibrils are made of proteins, which in turn form the thick filaments and the thin filaments.

As stated at the beginning, when striated muscle is examined under a microscope, a distinct repeating pattern of bands can be observed. Each repeating unit (also known as the sarcomere ), formed by the Z, I, A, H, and the M bands, is the product of the interaction between the proteins that form the thick filaments and the thin filaments. One of the goals of our lab is to characterize the proteins that form the sarcomere (sarcomeric proteins), hoping to identify new genes that are associated with disorders of muscle contraction.

Research has already unraveled many of the sarcomeric proteins that are responsible for skeletal muscle contraction. The names of some of these proteins are actin, myosin, troponin, tropomyosin, and nebulin. When skeletal muscle fibers receive a nerve impulse, these proteins (and others yet to be identified) are believed to change their shape and position. The thick filaments pull on and slide along the thin filaments, causing the sarcomere to shorten and the muscle to contract.

Just like the rest of the proteins in the body, instructions to make the sarcomeric proteins are encoded by different genes in the DNA . The DNA is actually a long string formed by four chemicals, namely, Adenine, Guanine, Cytosine, and Thymine (A, G, C, and T). The order in which these chemicals are found is what determines the genetic code, or sequence. A change in this order (i.e. mutation) results in a change of the encoded protein.

Proteins play many roles in the body. For example, they give the cues for development, determine eye color, and help in food digestion, among others.

Proteins play many roles in the body. For example, they give the cues for development, determine eye color, and help in food digestion, among others. Proteins are essential for body function and therefore, certain protein alterations can cause disease. In particular, alterations in any of the sarcomeric muscle proteins can potentially cause muscle disease. For example, alterations in actin are known to cause nemaline myopathy , a disease of skeletal muscle that causes low muscle tone, muscle weakness and respiratory difficulties, among others. Protein alterations are caused by gene changes ( mutations ).

At this point, it is worth mentioning that not all genetic changes have an impact on our health. Although the vast majority of the DNA is very similar among different people, some parts of the DNA may vary. All human beings are different. People have different skin color, hair, and eye color. The reason for these differences is that everybody's DNA is different. Most of the time, the DNA changes that produce different skin or eye colors do not have an impact on our health. In addition, many DNA changes are "silent", having no known effects on our development.

Silent" DNA changes do not cause disease or differences between individuals. When a new mutation is found in an individual, it is important to confirm whether the DNA change is disease causing or not. In the Beggs Laboratory , we do that by comparing samples from affected to non-affected family members, as well as to samples from members of the general population. If a mutation is present in both affected and non-affected family members, then we are likely to conclude that this mutation does not cause disease.

A genetic change that interferes with somebody's health must be significant enough so that the function of the protein encoded by the gene becomes altered. The outcome of a particular gene alteration is also dependent on how essential to the body is the protein made by that gene. Certain mutations in the genes encoding actin, nebulin, troponin, and tropomyosin cause nemaline myopathy , a disease associated with muscle weakness and respiratory problems. Similarly, mutations in the gene for myotubularin 1 cause X-linked myotubular myopathy , a severe condition associated with muscle weakness, skeletal problems, and fatigability. Specific genetic cause(s) for multiminicore disease, congenital fiber-type disproportion (CFTD), as well as certain non-specific myopathies are the subject of current research.

Identification of mutations may lead to better understanding of basic muscle biology, will allow for the development of improved diagnostic tests, and will hopefully lead to insights into therapies. This is why the Beggs Laboratory is actively looking for congenital myopathy-causing genetic changes.

The Role of Airway Smooth Muscle during an Attack of Asthma Simulated In Vitro

Excessive narrowing of airways in response to contractile agonists is a characteristic feature of asthma. We hypothesized that airway smooth muscle (ASM) adaptation to short lengths could contribute to exaggerated airway narrowing during an acute attack of asthma by allowing the muscle to regain its ability to generate maximal force at a shortened length. To test this hypothesis we mimicked, in vitro, the sequence of contractile events that would occur during a spontaneous attack of asthma. Trachealis muscle was challenged with carbachol (300 nM, submaximal dose) and allowed to shorten to approximately half of its original length. After 30 min of adaptation at the shortened length in the presence of carbachol, muscle force, amount and rate of shortening in response to electrical stimulation were compared with corresponding values obtained from control experiments during which the ASM was not adapted to the short length. After adaptation at the shortened length the developed force, amount and rate of shortening increased by 1.93 ± 0.08-, 1.57 ± 0.12-, and 1.75 ± 0.2-fold, respectively. Shortening of ASM in response to contractile agonists can lead to adaptation of the muscle to the shortened length that, in turn, can result in further shortening and the potential for airway closure.

Asthmatic airways in vivo narrow excessively in response to a number of stimuli, including dry air, hypo- and hypertonic aerosols, cholinergic agonists, histamine, and allergens (1–3). Excessive airway narrowing could be related to an increased amount of airway smooth muscle (ASM) (4–8), which would allow the development of greater wall stress and an increased ability to narrow against opposing loads. Alternatively, greater ASM shortening could be due to an alteration in muscle phenotype so that the same amount of muscle can generate greater force and shortening than normal ASM (9). However, most investigations of asthmatic ASM in vitro have not reported greater ASM force, active stress, or shortening than control ASM (for a review, see Ref. 10). An additional potential cause of exaggerated airway narrowing is adaptation of ASM to shortened length. The ability of ASM to generate force decreases when it is acutely shortened (11, 12). However, when a sufficient period of time is allowed for ASM to adapt, it exhibits a remarkable ability to generate the same maximal force over a very broad length range (13, 14). This “plasticity” or “length-adaptability” (15) of ASM has been attributed to evanescent formation or subtraction of contractile units in series and parallel (13, 16–18) and cytoskeletal remodeling (19, 20). If a similar phenomenon occurs in vivo, it could have a profound effect on airway mechanics during or following acute spontaneous attacks of asthma. Shortened ASM could regain its ability to generate force and shortening leading to a vicious cycle of progressive airway narrowing.

Length adaptability of ASM has been previously demonstrated by repeatedly stimulating the tissue at timed intervals under isometric conditions. This method of adaptation does not accurately represent the process of ASM adaptation that could occur in the airways of an asthmatic lung in vivo. For this reason, we investigated length adaptability after the shortening of porcine trachealis using a pharmacologic stimulus under conditions that mimic spontaneous in vivo bronchoconstriction.

Adaptation of trachealis to length change has been demonstrated previously by many independent investigators (13, 14, 16–21). This phenomenon, however, has never been unequivocally demonstrated in bronchial smooth muscle. In this study, we have included an important additional investigation showing that bronchial smooth muscle also adapts to length change and is able to regain much of its ability to generate force at short lengths.

Tracheas and the left lungs were removed from adult pigs immediately after the animals were killed by an overdose of sodium pentobarbital, using a procedure approved by the Animal Ethics Committee of the University of British Columbia. Tissues were stored in ice-cold carbogenated (5% CO2 in oxygen) Krebs-Henseleit solution (details on the tissue preparation and solution composition are provided in the online supplement). Tracheal and bronchial muscle strips were prepared by dissecting away the mucosal and adventitial layers to produce a smooth muscle layer for mechanical measurements.

All muscles were “equilibrated” before experimentation. This was performed by using 12 s supra-maximal electrical-field stimulations (EFS) set to occur at 5-min intervals until a stable maximal force was achieved. The muscle was stretched to a starting length with a resting tension just enough to keep the muscle strip taut. We defined this length as reference length (Lref). The Lref identified in this manner was usually near the transition region between the ascending-limb and the plateau of the length–force curve determined later in the experiment. For both bronchial and tracheal strips, the force–length data points were obtained sequentially from short to long lengths. Incremental length changes were performed 60 s before stimulation. This process continued until the cumulative length increase produced a passive force with a magnitude approximately equal that of active force. At this length (∼ 2 × Lref), the muscle was allowed to adapt and reach its maximal potential for force generation. After full adaptation at this long length, the muscle was released to a length near Lref and again allowed to adapt fully at that length.

The protocols used to simulate asthmatic bronchoconstriction are shown in Figures 1A and 1B

Figure 1. Asthma simulation protocol represented by time-lines (not to scale) for force and length of trachealis strips (six pairs) in response to a range of interventions. Tissues were paired, with one used to measure change in force (A), and the other for measuring change in velocity and amount of shortening (B). The letters below the time-line of A denote the following interventions: a, tissue adapted to Lref (“s” and bar denote electrical field stimulation [EFS] and stimulation duration). b, tissue length decreased to 0.5 Lref and immediately stimulated. c, d, and e, recovery of force in a three-cycle EFS-adaptation protocol at Lref. f, tissues were contracted to CCh (300 nM) and shortened against a load to permit shortening from Lref to ∼ 0.5 Lref for 30 min (intervention simulates adaptation of the muscle to a short length before the final asthma attack). g, bathing solution was changed three times to remove CCh from the tissue. (Note: after the first wash, the load was temporarily reduced to zero to prevent the tissue from being stretched). h, after an additional ∼ 7 min of being at 0.5 Lref the tissue was stimulated (EFS) at the same length to simulate an asthma attack. An increase in active force (FA) above the response observed for intervention “b”, as indicated by the broken force curve, would be interpreted as being the result of length adaptation leading to excessive shortening of the tissue. The time events indicated in B are as follows: b′, shortening of the tissue against a load 50% of FA at 0.5 Lref (the slope of the gray line indicates velocity of shortening) the broken time-line before intervention b′ indicates that interventions “a” to “e” have been performed. c, d, e, f, and g are the same as in A. h′, repeat of intervention “b′.” An increased shortening (indicated by the broken length curve) would be interpreted as being the result of length adaptation that could be responsible for the greater airway narrowing observed in asthma attack.

The protocol described in Figure 1B is exactly the same as that described in Figure 1A , except that at time b′ and h′ isotonic shortenings were induced (instead of isometric contractions in Figure 1A ). An isotonic load equivalent to half of the isometric force obtained at time b ( Figure 1A ) was used for the isotonic contraction at times b′ and h′ ( Figure 1B ). The rate and amount of shortening measured at time h′ was compared with those measured at time b′. Note that the stimulus, load, and initial muscle length at times b′ and h′ were all the same the difference again was in the history of length adaptation.

Curve fitting, statistics, and other data analyses can be found in the online supplement.

Figure 2. Partially length-adapted active (filled circles) and passive (filled triangles) forces with standard errors were plotted as fractions of the maximal active isometric force (Fmax) as a function of muscle length for (A) tracheal and (B) bronchial smooth muscle preparations. Data points (open circles) labeled “a” are fully adapted active forces obtained at a length near 2 × Lref. The corresponding passive forces (open triangles) are labeled “b.” After full adaptation at the long length, the muscles were shortened (in the relaxed state) to a length near Lref. Subsequent to the shortening, active force decreased from that indicated by “a” to that indicated by “c.” Adaptation of the muscles at the shortened length resulted in a substantial recovery of force (direction and magnitude of force recovery are indicated by the arrows). The short dashes besides the arrows indicate the level of force after each EFS during adaptation (see text for more description).

After full adaptation at 2 × Lref (with maximal isometric force indicated by “a”), muscle preparations were acutely shortened to Lref and immediately stimulated to obtain isometric force at Lref without adaptation. As a result, a large decrease in active force was found (from “a” to “c”). Upon adaptation at Lref, active forces of tracheal and bronchial preparations recovered substantially (with magnitude and direction indicated by the arrows). The short dashes beside the arrows indicate the levels of isometric force following each stimulation. The final adapted force at Lref was not different from that at 2 × Lref, for both preparations.

The time course of force recovery was very similar in both preparations ( Figure 3 )

Figure 3. Time course of active force recovery (as percent of maximal recovery) for porcine tracheal (open triangles) and bronchial (filled circles) smooth muscle preparations during adaptation at ∼ Lref. The same data, in different forms, were also plotted in Figure 2 (short dashes by the arrows). The solid lines indicate the curve-fits to all data points in each group using an exponential equation and a maximum recovery of 100% for tracheal (black line) and bronchial (gray line) tissues. The two groups of data are not statistically (ANOVA) different.

Figure 4. Ratios of post- versus pre-adaptation values of isometric force, amount of shortening, and velocity of shortening. The force ratio was obtained from measurements made at time point h ( Figure 1A ) divided by measurements made at time point b. The shortening and velocity ratios were obtained from measurements made at h′ ( Figure 1B ) divided by those at b′.

We have demonstrated in this study that both tracheal and bronchial smooth muscles are able to adapt to changes in muscle length and maintain their ability to generate maximal force over a large length range. This is the first time that the phenomenon of length adaptation has been documented for bronchial preparations. (Note that we have reported this phenomenon previously [22] along with a similar report from Stephens and coworkers [23] in the form of abstracts.) Previous studies of ASM length adaptability were all done using tracheal smooth muscle (13, 14, 16–21). These results thus allow us to use the model of length adaptation developed from tracheal smooth muscle to describe the similar behavior observed in bronchial smooth muscle. Since we have hypothesized that excessive airway narrowing in asthma is, at least partially, a consequence of length adaptation, it is important that we have established that length adaptation can occur in bronchial smooth muscle. The reason we most often use the trachealis preparation is that the muscle bundle contained relatively little connective tissue and therefore very little passive force is elicited when the bundle is stretched. This has allowed us to study the mechanical properties of smooth muscle cells with little “contamination” from the nonmuscle components of the preparation.

The major finding of this study is that ASM is able to adapt and regain its ability to generate maximal force and shortening after a period during which the muscle is submaximally activated and shortened. One important difference between the present finding and previous findings from studies of ASM length adaptation is that the present protocols include adaptation of the muscle using continuous stimulation by a contractile agonist (submaximal dose of carbachol). We believe that this protocol is a more accurate simulation of the in vivo environment of asthmatic airways, where smooth muscle is probably chronically exposed to submaximal levels of contractile agonists that initially may cause limited shortening but with progressive length adaptation, a subsequent additional stimulus is capable of causing enhanced shortening and narrowing of the airways, contributing to the asthmatic attack.

The increase in force due to length adaptation was associated with increases in both the amount and rate of shortening, against a constant load ( Figure 4 ). Since airway narrowing seen in asthma is closely linked to muscle shortening, the demonstration that the amount of shortening can be augmented through length adaptation allows us to interpret the present results directly in terms of airway narrowing.

The ability of ASM to adapt continuously while shortening (or in a statically shortened state) and regain its contractility, as demonstrated for the first time in this study, puts airways in a precarious state (in terms of their ability to maintain patency) when inflammatory mediators are present. There are many in vivo conditions under which ASM could adapt at abnormally short lengths (10, 24, 25). Chronic stimulation by inflammatory mediators could lead to muscle shortening over a prolonged period of time during which adaptation can occur. Remodeling of the airways as a result of chronic airway inflammation, which occurs in asthma and COPD, may also uncouple ASM cells from the periodic stretches imposed by tidal breathing and deep inspirations that have been shown to have bronchodilating effects (26–35). The decoupled ASM cells could adapt to a shortened length. A similar mechanism could be responsible for the airway hyperresponsiveness associated with sleep and paraplegia. During recumbent sleep, lung volume decreases and the unloaded ASM could adapt to the shortened length such a mechanism could be one of the contributors to the characteristic exacerbation of airway narrowing that occurs during sleep in asthma. A similar mechanism could contribute to the airway hyperresponsiveness that occurs after high spinal cord injury that lowers functional residual capacity and prevents full lung inflation (36).

In summary, these data suggest that length adaptation can alter the capacity of ASM to shorten. Adaptation to short length is one mechanism that could contribute to the exaggerated airway narrowing that is a feature of inflammatory airway diseases. A corollary of these results is the possibility that adaptation to long lengths could provide a beneficial bronchoprotective effect.

The authors thank the staff at the Animal Laboratory of the Jack Bell Research Centre for the supply of porcine tracheas.

Aims and scope

Skeletal Muscle is a peer-reviewed, open access, online journal that publishes articles investigating molecular mechanisms underlying the biology of skeletal muscle. A wide range of skeletal muscle biology is included: development, metabolism, the regulation of mass and function, aging, degeneration, dystrophy and regeneration. The emphasis is on understanding adult skeletal muscle, its maintenance, and its interactions with non-muscle cell types and regulatory modulators.

Peripheral Airway Smooth Muscle, but Not the Trachealis, Is Hypercontractile in an Equine Model of Asthma

Heaves is a naturally occurring equine disease that shares many similarities with human asthma, including reversible antigen-induced bronchoconstriction, airway inflammation, and remodeling. The purpose of this study was to determine whether the trachealis muscle is mechanically representative of the peripheral airway smooth muscle (ASM) in an equine model of asthma. Tracheal and peripheral ASM of heaves-affected horses under exacerbation, or under clinical remission of the disease, and control horses were dissected and freed of epithelium to measure unloaded shortening velocity (Vmax), stress (force/cross-sectional area), methacholine effective concentration at which 50% of the maximum response is obtained, and stiffness. Myofibrillar Mg 2+ -ATPase activity, actomyosin in vitro motility, and contractile protein expression were also measured. Horses with heaves had significantly greater Vmax and Mg 2+ -ATPase activity in peripheral airway but not in tracheal smooth muscle. In addition, a significant correlation was found between Vmax and the time elapsed since the end of the corticosteroid treatment for the peripheral airways in horses with heaves. Maximal stress and stiffness were greater in the peripheral airways of the horses under remission compared with controls and the horses under exacerbation, potentially due to remodeling. Actomyosin in vitro motility was not different between controls and horses with heaves. These data demonstrate that peripheral ASM is mechanically and biochemically altered in heaves, whereas the trachealis behaves as in control horses. It is therefore conceivable that the trachealis muscle may not be representative of the peripheral ASM in human asthma either, but this will require further investigation.

Our data show that the trachealis of horses with heaves behaves similarly to those of control horses, whereas the peripheral airway smooth muscle, which is more readily exposed to inflammation, is hypercontractile (increased unloaded shortening velocity). Moreover, we found a correlation between the muscle hypercontractility and the time elapsed since the end of the corticosteroid treatment.

Asthma is characterized by airway inflammation and airway hyperresponsiveness (AHR), an exaggerated bronchoconstrictive response to various stimuli (1). Because airway smooth muscle (ASM) is the direct effector of bronchoconstriction, it is believed to be hypercontractile in asthma. Indeed, several studies, performed in animals with experimentally induced or innate AHR, have shown that tracheal and bronchial smooth muscle (SM) is hypercontractile, exhibiting increases in maximal velocity of shortening (Vmax) (2–7) or maximal isometric force (4, 6, 8). To the contrary, measurements of these parameters in human tissues were rather inconclusive. For instance, in human trachealis and main bronchi muscle strips, isometric force (9–11) or Vmax (9, 10) were not different between subjects with asthma and control subjects. These results were corroborated by electron microscopy measurements showing that there were no structural differences between asthmatic and control trachealis (12). In contrast, stress and maximal extent of shortening was increased in bronchial strips of two patients with asthma (13, 14), although one of the subjects also suffered of lung carcinoma (14). In addition, in one study performed with fatal asthmatic trachealis samples, an increased isometric force and impaired relaxation were reported (11). Finally, bronchial spirals from fatal (11, 15) and nonfatal asthmatic lungs (16) showed hypocontractile responses.

There are several reasons why we may not clearly detect enhanced contractility in asthmatic ASM. The most obvious possibility is that the ASM is mechanically normal in asthma and that alterations occur at different levels. For example, the neural control could be altered in asthma (17) or airway–parenchymal interdependence could be reduced (18, 19), and these effects would not be seen in excised SM. Furthermore, whereas numerous studies point to an increased ASM mass (20), muscle force is always studied with normalization to cross-sectional area (CSA) so that this contribution is overseen. Another possibility is that the trachealis muscle, which is the muscle of choice to study ASM mechanics because of its ease of dissection, is not mechanically representative of the SM of more peripheral airways, which are likely to be involved in asthma. It is also possible that inflammatory mediators, present in greater amounts in the peripheral airways (21, 22), alter the ASM mechanical properties. Indeed, we recently showed that incubating Brown Norway rat trachealis muscle with CD4 + T cells for 24 hours increases its Vmax along with alterations of the contractile protein expression (7).

Thus, the purpose of this study was to compare the mechanical, biochemical, and structural properties of the trachealis and more peripheral ASM in horses with heaves, a naturally occurring model of asthma (23). This animal model offers a unique opportunity to dissect and study the intrapulmonary ASM mechanics. We report that the trachealis muscle is not mechanically different between heaves-affected horses and controls, whereas the SM of the peripheral airways is hypercontractile in heaves. Some of the results of this study have been previously reported in the form of an abstract (24).

Horses from the Faculty of Veterinary Medicine, University of Montreal (St.-Hyacinthe, PQ, Canada), were studied (see clinical details in Table 1). Animals were pooled into three groups according to their symptomatic history: (1) the control group contained horses with no history of respiratory diseases (2) the clinical remission group contained heaves-affected horses with inflammation and reversible airway obstruction upon hay exposure the remission state was obtained using antigen avoidance strategies (none of the horses had received corticosteroids for at least 4 months) and (3) the heaves group contained horses in clinical exacerbation of the disease (all of these horses except for one had also received some corticosteroid treatments) (see Table 1). All procedures were approved by the Animal Care Committee of the University of Montreal (Protocol Rech-1324) and complied with the guidelines of the Canadian Council on Animal Care.

Table 1. Clinical Information

Definition of abbreviation: NA, not applicable.

The animals were pooled into three main groups according to their symptomatic history: (1) the control group contained horses with no history of respiratory diseases (n = 5) (2) the clinical remission group contained heaves-affected horses with inflammation and reversible airway obstruction upon hay exposure (n = 3) the remission state was obtained using antigen-avoidance strategies (none of the horses had received corticosteroids for at least 4 months) and (3) the heaves group contained horses in clinical exacerbation of the disease (n = 5). Most of these horses had also received some corticosteroid treatments as described.

The lower part of trachea (10–15 rings) and intrapulmonary airways (3–7 mm inner diameter from degassed lungs) were placed in Hank’s balanced salt solution and dissected from connective tissues and parenchyma. Fine dissection (removal of the epithelium, cartilage, and connective tissues) was performed on ice in Ca 2+ -free Krebs-Henseleit solution. The buffers composition is detailed in the online supplement.

The muscle strip was mounted horizontally with foil clips to a length controller (no. 322C-I Aurora, ON, Canada) at one end and a force transducer (Aurora no. 400A) at the other, controlled by Aurora 600A software in a temperature controlled chamber (37°C) as described previously (7, 25). Reference length (Lref the in situ length under relaxed conditions) (9) and width at Lref were measured using a Hitachi camera (KP-D20A/B Hitachi, Toronto, ON, Canada) at 10–12 × magnification in Ca 2+ -free Krebs-Henseleit solution and analyzed with ImageJ software (IJ1.46r National Institute of Health, Bethesda, MD). Muscle strips from tracheal and intrapulmonary airways SM were used to study the methacholine (MCh) dose response, stress, stiffness, and shortening velocity. The details of the equilibration and mechanical measurements are included in the online supplement.

The total muscle strip CSA in square millimeters was calculated as previously described (26), with the details given in the online supplement. To evaluate the SM CSA, the muscle strips were processed for histology and examined by light microscopy, as detailed in the online supplement.

The in vitro motility assay was performed as previously described (27), and detailed in the online supplement.

The ATPase assay was performed as previously described (28), with modifications (29), and as detailed in the online supplement.

The Western blot was performed as previously described (7), and as detailed in the online supplement.

Data are presented as mean (±SEM). The n values refer to the number of animals from which two or three repeats were averaged. All parameters were analyzed using GraphPad Prism 5 (GraphPad Software Inc., La Jolla, CA) and MatLab (Natick, MA), as described previously (7). A two-way ANOVA (linear mixed model with airway location and disease stage as categorical variables) were performed, followed by a Bonferroni post hoc test. Correlation analysis was performed by GraphPad InStat version 3.10 using Pearson’s test. Differences were considered significant at a P value less than 0.05.

To assess whether the trachealis muscle is mechanically representative of the intrapulmonary ASM in heaves, we measured their mechanics in control and in heaves-affected horses during both clinical exacerbation and remission. There was a significant difference between groups with airway location (P = 0.01) and an interaction between airway location and stages of the disease (P = 0.018). No differences were observed between groups in Vmax of the trachealis muscle ( Figure 1A ). In contrast, Vmax was significantly greater in the peripheral ASM of heaves-affected horses in exacerbation (0.26 ± 0.05 Lref/s) compared with controls (0.12 ± 0.01 Lref/s P = 0.036), but not compared with the peripheral ASM of horses with heaves under remission (0.17 ± 0.02 Lref/s P = 0.21 Figure 1A ). Moreover, Vmax of the peripheral ASM of heaves-affected horses in exacerbation was significantly greater than that of their own trachealis (0.12 ± 0.01 Lref/s P = 0.036), control trachealis (0.11 ± 0.01 Lref/s P = 0.036), and the trachealis of horses under remission (0.12 ± 0.03 Lref/s P = 0.05 Figure 1A ). Importantly, in the heaves group, a linear correlation was observed between Vmax and the time elapsed since their last corticosteroid treatment for the peripheral airways (r 2 = 0.598 P = 0.009), but not for the tracheal SM (r 2 = 0.147, P = 0.27 Figure 1A , inset). Figure 1B shows representative force–velocity curves.

Figure 1. (A) Unloaded shortening velocity (Vmax) of tracheal and peripheral airway smooth muscle (ASM) from controls (open circles n = 5 horses), horses with heaves during exacerbation (solid circles n = 5 horses), and horses experiencing remission (crossed squares n = 3 horses) of the disease. The two circled values are from horses affected with heaves but recently treated with corticosteroids (horses 8 and 9 in Table 1). *P < 0.05. Inset, Vmax of tracheal (open circles) and peripheral (solid circles) ASM as a function of time between the last corticosteroid treatments and killing, measured in the horses with heaves. Lo and Lref, reference length. (B) Representative force–velocity curves for tracheal control (open diamonds) or peripheral control (open squares) smooth muscle (SM) and tracheal heaves–affected (solid diamonds) or peripheral heaves–affected (solid squares) SM. F/Fmax, force normalized to maximal force. The force–velocity relationships were accurately fitted by the Hill hyperbolic model (r 2 > 0.98). Data are presented as mean (±SEM).

MCh dose–response curves were performed for tracheal and peripheral ASM and are expressed in terms of stress ( Figures 2A–2C ) by normalizing by SM CSA ( Figure 2F ). For the maximal stress, there was no significant difference between airway location (P = 0.31), nor interactions with stages of the disease (P = 0.12). However, significant differences were observed between peripheral airways of the remission group and the control (P = 0.039) or heaves (P = 0.049) groups ( Figures 2C ). Similarly, the active stiffness was not different between airway location (P = 0.74), and there was no interaction with stages of disease (P = 0.10), but a significant difference was observed between peripheral airways of the remission group and control (P = 0.049) or heaves-affected (P = 0.049) animals ( Figure 2E ). The MCh dose at which 50% of the maximum stress is generated (EC50), was not different between groups no significant differences were found between airway location (P = 0.88) and stages of the disease (P = 0.83) ( Figure 2D ). Representative cumulative MCh dose–response traces of tracheal and peripheral ASM are shown in Figures 2G and 2H .

Figure 2. Mean cumulative methacholine (MCh) dose–response curves expressed as SM stress for (A) tracheal SM and (B) peripheral ASM from control horses (open circles n = 4 horses), heaves-affected horses (solid circles n = 5 horses), and horses under remission (open squares n = 3 horses). (C) Maximal stress generated by tracheal and peripheral ASM at 10 −4 M MCh (maximal force normalized to SM cross-sectional area [CSA]) in control (white bars), heaves (black bars), and heaves under remission (gray bars). *P < 0.05. (D) Effective concentration at which 50% of the maximum stress (EC50) is generated for the three groups of animals. (E) Active stiffness (force normalized to SM CSA and divided by Lref) measured in tracheal and peripheral ASM of control horses (white bars), horses with heaves (black bars), and horses under clinical remission (gray bars). *P < 0.05. (F) Cross-section of tracheal and peripheral ASM strips used for CSA quantification. (G and H) Representative cumulative MCh dose–response traces of tracheal (G) and peripheral airway (H) SM. Arrows denote time points of MCh injection. Data are presented as mean (±SEM).

To study whether the biochemical properties were altered in tracheal and peripheral ASM of control and heaves-affected horses, measurements of myofibril Mg 2+ -ATPase activity were performed. Isolated myofibrils showed significant differences in Mg 2+ -ATPase between airway location (P = 0.015) and the interaction between airway location and stages of the disease (P = 0.013). Myofibrils isolated from peripheral ASM of heaves-affected horses showed significantly higher ATPase activity (117 ± 24.8 nmol inorganic phosphate (Pi)/mg actomyosin (AM)/min) than myofibrils isolated from control peripheral ASM (28.5 ± 14.9 P = 0.019), control trachealis (30 ± 14.1 P = 0.02), and heaves-affected horses trachealis (33.9 ± 12.8 P = 0.026) ( Figure 3 ).

Figure 3. Mg 2+ -ATPase activity of myofibrils isolated from tracheal and peripheral ASM of control (open circles and open squares, respectively n = 3 horses for both) and heaves-affected horses during exacerbation (solid circles and solid squares n = 3 horses for both). *P < 0.05. Data are presented as mean (±SEM).

To verify whether or not the molecular mechanics of the myosin motor is also altered in the peripheral ASM of the heaves-affected horses, we purified myosin from tracheal and peripheral ASM and measured the velocity of actin filament propulsion (νmax) in the in vitro motility assay. There was a significant difference between airway location (P = 0.0002), but not at the interaction between airway location and stages of the disease (P = 0.66). No differences in νmax were observed between control (0.48 ± 0.05 µm/s) and heaves under exacerbation (0.49 ± 0.04 P = 0.84) of tracheal SM or between control (0.3 ± 0.03 µm/s) and heaves under exacerbation (0.29 ± 0.01 P = 0.72 Figure 4 ) of peripheral ASM. Note that νmax of myosin purified from control tracheal SM was significantly greater than that of peripheral ASM of controls (P = 0.002) and heaves-affected horses (P = 0.003), and that νmax of myosin from tracheal SM of heaves-affected horses was significantly greater than that of peripheral ASM of control (P = 0.001) and heaves-affected horses (P = 0.001).

Figure 4. Velocity of actin filament (νmax) when propelled by thiophosphorylated myosin purified from tracheal and peripheral ASM of controls (open circles and open squares, respectively n = 3 horses for both) or heaves-affected horses (solid circles and solid squares n = 5 horses for both), as measured in the in vitro motility assay. Arrows show νmax for myosin purified from tracheal or peripheral ASM of horses affected by heaves recently treated with corticosteroids. *P < 0.0001. Data are presented as mean (±SEM).

To determine whether the differences in peripheral ASM mechanics were due to alterations in contractile protein expression, the level of the (+)insert SM myosin heavy chain (SMMHC) isoform (or SMB), calponin (CaP), total SMMHC, myosin light chain kinase (MLCK), and transgelin (SM22) was quantified ( Figure 5 ). No differences between airway location or stage of disease were seen in the expression of SMB (P = 0.34 P = 0.077 Figure 5A ), total SMMHC (P = 0.75 P = 0.93 Figure 5C ), and SM22 (P = 0.1 P = 0.18 Figure 5E ). Significant differences were measured for CaP ( Figure 5B ) and MLCK ( Figure 5D ) between airway location (P = 0.008 and P = 0.0003), but not between stages of disease (P = 0.2 and P = 0.66, respectively). CaP in the tracheal SM of heaves-affected horses was significantly greater (2.15 ± 0.57 arbitrary units) than in peripheral ASM of controls (0.37 ± 0.14 P = 0.033) and heaves-affected horses (0.4 ± 0.16 P = 0.026 Figure 5B ). The expression of MLCK in control trachealis muscle (0.84 ± 0.13) was greater than that of control peripheral ASM (0.05 ± 0.003 P = 0.03). Similarly, the expression of MLCK in heaves-affected horses trachealis muscle (0.83 ± 0.24) was greater than that of control peripheral ASM (0.05 ± 0.003 P = 0.024 Figure 5D ).

Figure 5. Western blot analysis: (A) (+) insert SM myosin heavy chain (SMMHC) isoform B (SM-B) (B) calponin (CaP) (C) total SMMHC (D) myosin light chain kinase (MLCK) and (E) transgelin (SM22) of tracheal SM (TSM) of control (open circles) or heaves-affected horses (solid circles) and peripheral ASM (PASM) of control (open squares) or heaves-affected horses (solid squares) n = 4 for TSM and PASM of control horses (except for SM22, where n = 3 horses) n = 5 for TSM and PASM of heaves-affected horses (except for SM22, where n = 4 horses). Smooth muscle actin (smA) was used as a loading control. The circled values are from horses affected with heaves that were recently treated with corticosteroids (horses 8 and 9 in Table 1). Representative Western blots are shown. *P < 0.05. Data are presented as mean (±SEM).

In this study, we compared the contractile and biochemical properties of the tracheal and peripheral ASM in heaves, a spontaneously occurring asthma-like disease in horses.

Very few studies have investigated the force generation and velocity of shortening of human asthmatic ASM because of the difficulty in obtaining such human samples. Those few studies, however, did not reveal the expected enhancement in contractility. To the contrary, the results were rather inconclusive. Some studies reported increased contractility in human asthmatic trachealis and bronchi strips or spirals (11, 13,14) or in sensitized bronchi (30), whereas others reported no mechanical differences between human asthmatic and control tracheal and main bronchi muscle strips (9, 10) or bronchial spirals (11, 15, 16). One of the problems of the early studies is that the tissues were often obtained several hours postmortem. Conversely, the more recent studies have used transplant-grade lungs, but have addressed only the mechanical properties of the trachealis and main bronchi SM (9, 10), and not those of peripheral ASM because of the difficulty of dissection. If the ASM is not intrinsically altered in asthma, but its contractility is only enhanced when exposed to inflammatory cells and mediators, then the peripheral airways, which are more exposed to inflammation (21, 22), are more likely to have hypercontractile SM. Indeed, we recently demonstrated in the Brown Norway rat model of asthma that ex vivo exposure of the trachealis muscle to inflammatory cells for 24 hours enhances its mechanical properties (7). Thus, in the current study, we addressed whether or not the trachealis SM behaves differently from the peripheral ASM in a species in which we can easily access and dissect the peripheral ASM.

Horses with heaves (23) develop moderate to severe airway obstruction when kept in an antigen-rich environment characterized clinically by labored breathing at rest (dyspnea). Treatment with corticosteroids, or prolonged antigen avoidance, lead to marked improvement of clinical signs and lung function (clinical remission of the disease) (31). Antigen avoidance was the strategy used in the current study to create clinical remission. Note that, because of the limitations in tissue availability, the muscle mechanics measurements were performed on all tissues, whereas the in vitro motility, ATPase activity measurements, and Western blots were performed only on subgroups (the n is indicated in the figure legends). Furthermore, the group under remission contained only three animals. This is a limitation of our study and, while it prevents us from drawing definitive conclusions regarding peripheral ASM mechanics in heaves, it generates hypotheses to be tested in human asthma.

Vmax measured in the trachealis muscle was not different between controls, horses with heaves, and horses with heaves under remission ( Figure 1 ). This is in agreement with the recent data obtained in human trachealis (9, 10). However, one of the most prominent results of our study was the roughly twofold-greater Vmax measured in the peripheral ASM of the horses with heaves compared with the control peripheral ASM and with all other trachealis SM ( Figure 1 ). In addition, Vmax of the peripheral airways of the horses with heaves under remission tended to decrease back toward baseline values, but this did not reach significance most probably because of the large variability in the heaves group. Two of the horses with heaves (the two circled values in Figure 1 ) had recently received corticosteroids and showed a somewhat lower Vmax. Horse no. 8 developed very pronounced heaves symptoms and had to be treated until the scheduled killing for ethical reasons. Although its symptoms improved, it still had a high pulmonary resistance (RL) (see Table 1) and was clearly still under exacerbation. The second horse (#9) that needed treatment with corticosteroids had developed lameness. This horse did not have very pronounced exacerbations and after the treatment transiently had a RL considered normal. Treatments with corticosteroids improve heaves symptoms, but this is only transient under continued antigen exposure. Thus, although it is obvious that the treatments received improved lung function and potentially contributed to decreasing their Vmax, the treated horses were not in a controlled state to fit the remission group. Furthermore, all of the horses with heaves except for one had received corticosteroids at given points in time. Conversely, horses with heaves under remission had not been antigen exposed and had not exhibited airway obstruction for at least 3 months, and had not received corticosteroids for at least 4 months.

A strong correlation between the time elapsed since the last corticosteroid treatment and Vmax of peripheral ASM was found in the horses with heaves under exacerbation ( Figure 1A , inset). The mechanism behind this effect might be related to a decrease in specific inflammatory cells or mediators in peripheral airways of the heaves-affected horses during treatment. However, it has been shown previously, in a similar cohort of horses, that inhaled corticosteroids do not significantly decrease the number of neutrophils, lymphocytes, mast cells, and macrophages during the first month of treatment (31). Thus, either the observed effect occurred via another type of cell/mediator combination or it is a direct effect of the corticosteroids on the muscle (32).

There are no differences in dose responses between controls and the animals with heaves, either in terms of maximum stress or effective concentration at which 50% of the maximum response is obtained for both the trachealis and the peripheral airways ( Figure 2 ). This is in agreement with the recent stress data obtained in human trachealis in response to electrical field stimulation (EFS) (9) or MCh (10) stimulation. However, we observed significant differences in stress and stiffness of peripheral ASM between the remission and the control groups or remission and heaves under exacerbation ( Figure 2E ), potentially due to airway remodeling. Several studies have addressed how ASM changes its stiffness and contractility after mechanical oscillation and stretch, because this is believed to be an important contributor to airway normoresponsiveness (9, 33), but we did not test for such effects on muscle contractility.

Intuitively, one would expect that ASM force generation would be increased in AHR and asthma/heaves, but not the velocity of shortening. However, a greater Vmax was also reported in the canine ragweed allergic model of AHR (3) and in the Brown Norway rat ASM exposed to CD4 + T cells (7). A stronger ASM would lead to a more pronounced bronchoconstriction upon simulation, but this has yet to be demonstrated convincingly in asthmatic tissues (9, 13). Conversely, a faster ASM could lead to greater airway resistance by counteracting the relaxing effect of tidal breathing (34). Extensive studies have shown that tidal or deep breaths stretch ASM and decrease airway resistance in normal lungs, but not in patients with asthma (35). Furthermore, using imaging techniques, Brown and coworkers (36) have shown that asthmatic airways dilate upon stretching, but that this is transient, because they rapidly narrow back to their initial diameter. This suggests that asthmatic ASM can shorten faster than normal ASM after a stretch. Thus, this “fast” muscle might be responsible for maintaining asthmatic airways in a more constricted state, because the ASM has time to shorten significantly between each breath, thus counteracting the relaxing effect of tidal and deep breaths.

To further dissect the mechanisms that led to the increased Vmax in the airways of heaves-affected horses, we investigated the biochemical activity of these SMs at the myofibrillar level. Previous studies have shown that Vmax is proportional to the rate of ATP hydrolysis in skeletal and SM (37). Indeed, supporting our muscle mechanics results ( Figure 1 ), the myofibrillar Mg 2+ -ATPase activity was roughly threefold greater in the peripheral airways of the heaves-affected horses compared with controls and to the trachealis muscle of heaves-affected horses ( Figure 3 ). Similar results were obtained in other animal models however, in those studies, Vmax (2, 3) and ATPase activity (3, 38) were increased in both the tracheal and peripheral ASM. These differences may be attributable to the mode of sensitization or to different asthma animal models. The Fisher and Lewis rat model of innate AHR (8) also showed a greater isometric tension for the trachealis of the hyperresponsive Fisher rats, which suggests that there might be cases of intrinsic alterations of the ASM. Thus, the similarities in tracheal Vmax and stress between controls and horses with heaves suggest that the equine model is mechanically more representative of human asthma.

To investigate whether the difference in Vmax and ATPase activity can be attributed, at least partially, to the mechanics of the myosin molecules, we purified the ASM myosin, the molecular motor of muscle contraction, and measured its νmax of actin propulsion in the in vitro motility assay. No differences were observed in νmax between control and heaves-affected horses ( Figure 4 ). Similar results were obtained with tracheal SM myosin purified from the ragweed-sensitized dog model (39), despite an increased Vmax (3) and ATPase activity (38) in that model. This suggests that myosin alone is not sufficient to alter Vmax, but that other proteins, or post-translational alterations that might have been eliminated during the myosin purification procedure, are required. Finally, it is noteworthy that νmax for the peripheral airways was lower than that of the trachea, both in controls and in horses with heaves, most probably because less SM is available for the purification in the peripheral airways, which might affect the quality of the resulting myosin.

In the current study, the Western blot (WB) analysis did not show major differences between controls and the heaves groups ( Figure 5 ). The choice of the contractile proteins analyzed by WB was based on our previous study (40) that examined the messenger RNA expression in endobronchial biopsies from normal subjects and patients with asthma in which we observed a greater expression of the (+)insert SMMHC isoform, MLCK, and SM22 in the patients with asthma. More (+)insert SMMHC and MLCK pointed toward a more contractile phenotype, whereas the function of SM22 remains unknown. We did not observe any significant differences in the fast (+)insert SMMHC isoform expression in tracheal and peripheral ASM of control and heaves-affected horses. Accordingly, no differences were observed in νmax in the motility assay ( Figure 4 ). Note that, in a previous study, we reported a greater expression of the fast (+)insert SMHHC isoform in the peripheral airways of the heaves-affected horses compared with peripheral airways of controls and horses with heaves under remission (41), but variability between animals and airway location may account for the differences between studies along with the different technology used to quantify protein content (Western blot versus mass spectrometry). The expression of MLCK and CaP showed significant differences between airway locations, but not between disease stages. These results are difficult to interpret, because expression of these proteins may also have been influenced by the corticosteroid treatments ( Figures 5B and 5D ). It is interesting to note that a lower MLCK content in airways was observed in the ragweed-sensitized dog model, where the activity of MLCK, but not its content was elevated (42).

One surprising result from WB analysis was the significantly greater CaP expression in trachea of heaves-affected horses compared with peripheral ASM of control and heaves animals ( Figure 5 ). It is unclear why CaP should be expressed differently in the trachealis and not in the peripheral airways of the heaves-affected horses. CaP inhibits myosin cycling through its interaction with F-actin. Indeed, it has been shown at the tissue level that Vmax of CaP knockout mice is higher than that of wild-type mice, and that adding back exogenous CaP inhibits the unloaded Vmax (43, 44). We obtained additional data comparing the contractile protein profile of the myofibrils described previously here, and our results suggest that filamin and α-actinin are also differently expressed between the trachea and the peripheral airways and potentially differently expressed between heaves and control trachealis (on line supplement). These proteins are known inhibitors of the Mg 2+ -ATPase activity, νmax and Vmax (45, 46), and this could have contributed to the decreased Vmax and Mg 2+ -ATPase activity that we observed in tracheal SM relative to peripheral SM from horses with heaves under exacerbation.

It is conceivable that several factors are required for a muscle to become hypercontractile. For example, the presence of inflammatory cells and mediators, or the possibility to repetitively shorten excessively upon stimulation, might be necessary for a muscle to become hypercontractile. Mechanical adaptation and plasticity have indeed been studied extensively and are believed to contribute significantly to AHR (47, 48). Thus, the trachealis of the heaves-affected horse might be exposed, although to a smaller extent than the peripheral airways, to inflammation, but is restrained from excessive contraction due to the cartilage rings this may lead to a different muscle phenotype in which contractile proteins that favor muscle relaxation are up-regulated. This will obviously require further investigations.

The principal site of airway obstruction in asthma has been a topic of debate for several years. Whereas the peripheral airways have initially been termed “silent,” because their small size was compensated by the large total CSA of the peripheral airways (49), more recent studies have revealed several reasons why they may play an important role in asthma (50, 51). Our study did not address precisely which airway generation is involved in asthma, but assessed the function of extra- versus intrapulmonary airways. Thus, our observations could be interpreted in terms of the effects of intrapulmonary inflammation (21, 22). Nevertheless, even if it is well accepted that the trachealis and main bronchi SM are not involved in asthma, because of the restriction of their contraction due to the cartilage rings, they have been used extensively in animal models and in human studies to address AHR. Using the SM of those extrapulmonary airways to study asthma assumes that intrinsic mechanical properties of the ASM are altered. Our data demonstrate that this is not the case in the heaves model of asthma. Indeed, the trachealis of the heaves-affected animals behaves similarly to that of the controls, whereas the peripheral ASM exhibits hypercontractility. Thus, the peripheral ASM potentially acquires these abnormal mechanical properties in the presence of the inflammatory mediators (7) or, as suggested by others, because of repeated contractile challenges and adaptation of the SM to shorter lengths (47, 48, 52). The increased enzymatic activity that leads to an increase in Vmax appears to be the primary response of the ASM in this horse model of asthma however, the precise pathway involved will await further studies.

Our data show that the trachealis of the horses with heaves behaves similarly to that of control horses, whereas the peripheral ASM, which is more readily exposed to inflammation, is hypercontractile (increased Vmax). Having previously demonstrated that incubation of ASM with inflammatory cells enhances their mechanics (7), we conclude that the alterations in peripheral ASM mechanics of the horse with heaves must be induced by the inflammatory cells present in the peripheral airways.


Clinical samples

The subjects were 57 patients presenting with atherosclerosis plaques at Tianjin Chest Hospital between January 1 and December 31, 2018, and 57 healthy volunteers recruited as controls. Blood samples (3 ml) were collected from each subject. After the blood was centrifuged at 3000 g for 10 min, the plasma (no blood cells) was collected and stored at − 80 °C. All patients and volunteers signed informed consent and the research protocol was approved by the Ethics Committee of Tianjin Chest Hospital.

Animal model

The animal experiment was approved by the Animal Ethics Committee of Tianjin Chest Hospital. This experiment involved 60 C57BL/6 8-week old mice from the Department of Laboratory Animal Science of Peking University Medical School. Of these, 50 were apolipoprotein E-knockout mice (ApoE −/− mice) fed with high-fat diet for 12 weeks to induce atherosclerosis. The remaining 10 were wild-type mice fed a normal diet and used as the empty control.

For the experiment, 40 ApoE −/− mice were divided into four groups of 10: the miR-637 agomir negative control (AG-NC), miR-637 agomir (AG), miR-637 antagomir negative control (AN- NC) and miR-637 antagomir (AN) groups. One week after the mice had adapted to their environment, miR-637 agomir or miR-637 antagomir were dissolved into 0.2 ml saline at a dose of 20 mg/kg body weight/day, and respectively injected into the AG and AN group mice through the tail vein once every 2 weeks. After the end of the experimental period, the mice were intraperitoneally injected with 20% uratan at a dose of 7 ml/kg body weight as a general anesthetic. When the muscle tension of the mice was weakened and the breathing was accelerated, blood was carefully collected and stored in a 5 ml EP tube. Then, the blood samples were centrifuged at 3000 g for 10 min at 4 °C for 5 min to obtain the plasma. In addition, the common carotid artery of the mice was isolated under aseptic conditions. After being washed with PBS, the intima was gently scraped off and the outer membrane was removed. The vascular smooth muscle layer was retained for subsequent experiments.

Cell culture and transfection

Human VSMCs from the ATCC were cultured in RPMI-1640 containing 10% fetal bovine serum (FBS), 1% penicillin and 1% streptomycin (Gibco) at 37 °C and in 5% CO2. They were treated with different concentrations of ox-LDL (0, 25, 50 and 100 mg/l) for 24 h or with 100 mg/l ox-LDL for different times (0, 12, 24, 48 h). VSMCs in the log phase were selected for transfection. MiR-637 mimics and inhibitors were transfected using Lipofectamine 2000 (Invitrogen). Forty-eight hours later, the transfection efficiency was validated using real-time PCR.

RNA extraction and quantitative real-time PCR analysis

To confirm that the plasma samples were hemolysis-free, their hemoglobin levels were measured using spectrophotometry (Thermo Fisher Scientific) and quantified with the formula CHb = 1.58A415–0.95A450–2.91A700 at the threshold sensitivity of 0.01 mg/ml. All the plasma samples included in the study contained ≤0.2 mg/ml hemoglobin.

Then, real-time PCR was used to determine the relative expression of miR-637 and IGF-2 mRNA. Total RNA was extracted from atherosclerosis patient plasma, healthy volunteer plasma, C57BL/6 mouse plasma and human VSMCs with TRIzol reagent (Invitrogen) according to the manufacturer’s instructions. To detect the relative expression of IGF-2 mRNA, reverse transcription was conducted using MMLV transcriptase (Invitrogen) to generate cDNA. To detect the relative expression of miR-637, reverse transcription was performed with a TaqMan MicroRNA Reverse Transcription Kit (Thermo Fisher Scientific). Quantitative real-time PCR was performed on the ABI 7500 Real-Time PCR System (Applied Biosystems) with SYBR premix EX TAQ II kit (TaKaRa) according to the manufacturer’s instructions. GAPDH and U6 were used as internal controls. The relative expressions were determined using the 2 -ΔΔCt method. In this study, primer sequences were constructed with the aid of Primer Premier 6.0 and are shown in Table 1.

CCK-8 assay

VSMCs at the log phase were selected and seeded in a 96-well plate (5 × 10 3 cells/well) with 3 replicate wells for each sample type. Ten microliter CCK-8 solution (Hubei Baiaosi Bioscience) was added to each well. The OD value was measured at 450 nm using a microplate reader (Bio-Rad Laboratories).

Apoptosis assay

The cells were seeded into a 6-well plate. When the cells had grown to 60–70% confluence, the medium was discarded. The plate was washed twice with PBS solution, and cells were trypsinized with 0.25% trypsin. Then the cells were collected in an EP tube and centrifuged at 1000 r/min and 4 °C for 5 min. The supernatant was removed and the cells were resuspended with 500 μl binding buffer. An FITC Annexin V cell apoptosis assay kit (Ruibo) was used to stain cells for 30 min at room temperature. After that, the apoptosis level was determined using a flow cytometer (Becton Dickinson).

Transwell assay

The cells were serum-starved for 24 h and then trypsinized. The medium was discarded, the cells were washed twice in PBS solution, and then resuspended in serum-free medium containing BSA to adjust the cell density to 2 × 10 5 cells/ml. Afterwards, 200 μl of cell suspension was added to a Transwell chamber (8 μm pore size Corning), and the lower chamber was supplemented with RPMI-1640 containing 20% FBS. Following 24 h of culture at 37 °C and in 5% CO2, the culture medium in the upper chamber was discarded and the cells that had not passed through the membrane in the upper chamber were carefully wiped with cotton swabs. The upper chamber was then washed twice with PBS. Residual cells were fixed with 4% paraformaldehyde for 15 min and stained in 0.1% crystal violet for 30 min. Five fields were randomly selected under the microscope for counting.

Western blotting

The cells were lysed with RIPA solution (Beyotime Biotechnology) containing the protease inhibitor PMSF and the total protein was extracted. The protein concentration was determined using the BCA method. Equivalent amounts of protein were separated via 10% SDS-PAGE and transferred onto nitrocellulose membrane. After being blocked with 5% skim milk, the membrane was incubated with primary antibody at room temperature for 1 h. The membrane was then washed with TBST solution, and incubated with secondary antibody at room temperature for 1 h, followed by washing with TBST solution. After the addition of ECL solution (Hubei Baiaosi Bioscience), color development was carried out, and the protein bands were quantitatively analyzed using Image J.

Dual luciferase reporter assay

StarBase and TargetScan were used to predict the target sequence of miR-637. Wild-type (WT) and mutant-type (MUT) IGF-2 were subcloned into pGL3 vector (Promega) and transfected into VSMCs. The cells were then plated in 24-well plates with 1 × 10 5 cells/well. Then, luciferase activity was determined with the dual luciferase system (Promega). MiR-637 mimics were transfected into VSMCs using wild or mutant vector. Luciferase activity was measured 48 h after the transfection.

Statistical analysis

Statistical analysis was performed using SPSS 22.0. The measurement data are presented as means ± standard deviation. The independent samples t test was performed for the comparison between the two groups. The difference was statistically significant at p < 0.05.

Coupling Excitation to Contraction

  • In resting muscle fibers, Ca 2+ is stored in the endoplasmic (sarcoplasmic) reticulum.
  • Spaced along the plasma membrane (sarcolemma) of the muscle fiber are inpocketings of the membrane that form "T-tubules". These tubules plunge repeatedly into the interior of the fiber.
  • The T-tubules terminate near the calcium-filled sacs of the sarcoplasmic reticulum.
  • Each action potential created at the neuromuscular junction sweeps quickly along the sarcolemma and is carried into the T-tubules.
  • The arrival of the action potential at the ends of the T-tubules triggers the release of Ca 2+ .
  • The Ca 2+ diffuses among the thick and thin filaments where it
  • binds to troponin on the thin filaments.
  • This turns on the interaction between actin and myosin and the sarcomere contracts.
  • Because of the speed of the action potential (milliseconds), the action potential arrives virtually simultaneously at the ends of all the T-tubules, ensuring that all sarcomeres contract in unison.
  • When the process is over, the calcium is pumped back into the sarcoplasmic reticulum using a Ca 2+ ATPase.

We believe serious diseases deserve serious attention

Cytokinetics is a late-stage biopharmaceutical company focused on discovering, developing, and commercializing first-in-class muscle activators and best-in-class muscle inhibitors as potential treatments for people with debilitating diseases in which muscle performance is compromised and/or declining. A leader in muscle biology research with more than 60 publications, over 50 clinical trials, and hundreds of issued patents, Cytokinetics is developing small molecule drug candidates specifically engineered to impact muscle function and contractility.

Cytokinetics was founded in 1998 by pioneers in the field of muscle biology. Over the years, our company has developed an unparalleled expertise, keeping it at the forefront of drug discovery and development for diseases impacting muscle performance. Muscle plays a critical role in everyday physical functionality, including proper circulation, movement, and even the ability to breathe. As a result, conditions or syndromes that lead to dysfunction of muscle have serious consequences on survival and well-being.

Cytokinetics takes a purpose-driven approach by leveraging our unique muscle biology expertise to engineer compounds with specific characteristics aimed at treating diseases that impact muscle function. By directly targeting the sarcomere, the foundation of muscle contraction, the treatments we are developing have the potential to preserve and extend independence and self-reliance in people suffering from debilitating diseases. Cytokinetics is dedicated to helping underserved patient populations which lack effective therapies.

Cytokinetics scientists have discovered a robust pipeline of small molecule muscle activators and inhibitors omecamtiv mecarbil, a cardiac myosin activator reldesemtiv, a fast skeletal muscle troponin activator (FSTA) CK-274 and CK-271, cardiac myosin inhibitors CK-136, a cardiac troponin activator and CK-601, a next-generation FSTA. Our muscle biology drug discovery and development platform has great potential for many other diseases and medical conditions characterized by muscle weakness, fatigue, or diminished muscle function.

Watch the video: Cardiac Muscle Physiology Animation (October 2022).