Study Guide for Chapters 19, 20, and 21
Study Guide for Chapters 19, 20, and 21 BIL360
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1 Control of Movement: The Motor Bases of the Animal Behavior Chapter 19 A muscle spindle is an example of a proprioceptor—a mechanosensory receptor that is associated with the musculoskeletal system. Proprioceptors are important for the control of movement because they provide an animal with information about where the parts of its body are positioned in space—information that is necessary both to program a movement and to monitor how the movement is progressing. The muscle spindle organ monitors the length of a skeletal muscle. Vertebrate muscle-spindle stretch receptors (1) Skeletal muscles contain muscle spindles arranged in parallel with tension-producing extrafusal muscle fibers. The cell bodies of the 1a afferent sensory neurons are in the dorsal root ganglia. (2) Muscle spindles are sensitive to changes in muscle length. Each muscle spindle consists of eight to ten specialized muscle fibers, called intrafusal fibers, that are contained within a capsule of connective tissue (Part 1 of figure). Intrafusal fibers are in contrast to extrafusal fibers, which are the normal “working” contractile fibers that bear loads and generate movements. When an intrafusal fiber is stretched by elongation of the whole muscle, the sensory neuron terminal associated with the central portion of the fiber is distorted; the distortion increases the cation permeability of the nerve terminal membrane and produces a graded receptor potential which, if threshold is reached, gives rise to action potentials (Part 2 of figure). The functional organization of a muscle spindle has several salient features: 1. The intrafusal fibers of muscle spindles are too few and weak to generate appreciable tension themselves. Because they are located in parallel to the tension-producing extrafusal fibers, muscle spindles do not sense muscle tension. 2. Muscle spindles provide information about muscle length and, in some animals, about changes in length. Their activity thus provides information about both muscle length (position) and rate of change in length (velocity of movement). 3. The intrafusal fibers of muscle spindles in vertebrates receive motor innervation. In birds and mammals, a separate class of small motor neurons termed gamma (γ) motor 2 neurons makes synaptic contact on the contractile portions of the intrafusal fibers. Signals from the CNS that cause contraction of the ends of the intrafusal fibers regulate the degree of stretch on the central non-contractile region, and thereby influence the degree of distortion of the 1a afferent nerve terminal. The combined actions of alpha (α) motor neurons (to the extrafusal fibers), γ motor neurons (to the intrafusal fibers), and 1a afferent sensory information contribute to smooth, coordinated motions. Vertebrates have additional types of proprioceptors. Two examples are Golgi tendon organs, which are embedded in tendons in series with working muscle fibers and monitor muscle tension, and joint receptors, which are embedded in connective tissue at a joint and respond over a certain range of joint positions or movements. Spinal reflexes are mediated by the neural circuits of the vertebrate spinal cord. As a result of pioneering studies by Charles Sherrington (1857–1952) and Ivan Pavlov (1849– 1936) on spinal reflexes at the beginning of the twentieth century, analysis of behavior in terms of spinal reflexes dominated studies of neural circuits through most of the twentieth century. These spinal reflexes are therefore perhaps the best-known vertebrate neural circuits. In spinal reflexes, sensory input (from receptors of the skin, muscles, tendons, and joints) enters the spinal cord through the dorsal roots. This sensory input, via intervening synapses in the spinal cord, excites some motor neurons and inhibits others, leading to movements by selectively activating muscular contraction. The sensory inputs from different populations of receptors have different connections in the spinal cord and thereby initiate different reflexes. We will examine two of the many reflexes of the mammalian hindlimb that have been studied extensively: the stretch reflex and the flexion reflex. Description of the figure: Stretching of muscle spindle causes 1a afferent neurons to generate Aps. 1a input generates EPSPs in motor neurons; stretched muscle (extensor) contracts. 1a afferent axons also synapse with inhibitory interneurons that inhibit motor 3 neurons to antagonist (opposing) muscles (flexor). Stretching of a leg extensor muscle activates a muscle spindle stretch receptor. The sensory neuron of the stretch receptor synapses on motor neurons (E) to the same muscle. Motor neurons action potentials excite extensor muscle fibers, causing contraction. The stretch receptor sensory neuron also activates inhibitory interneurons that inhibit the motor neurons (F) to antagonistic muscle fibers. Reciprocity: muscles are arranged in antagonist pairs and signals that activate movements contract agonists while relaxing antagonists. THE FLEXION REFLEX When you step on a tack, you reflexively withdraw your foot from the offending stimulus. Your foot is drawn upward by contraction of the flexor muscles of the thigh. The neural circuit mediating this flexion reflex is shown at the left in Figure 19.2. A diverse array of sensory neurons known as flexion-reflex afferents have endings in the skin, muscles, and joints; some of these are sensitive to painful and noxious stimuli. The flexion-reflex afferents make excitatory synaptic contacts on interneurons in the CNS that in turn excite motor neurons to the flexor muscles, as well as inhibitory interneurons that inhibit motor neurons to the extensor muscles. Thus, as in the stretch reflex (and in other spinal reflexes), synaptic interactions in the spinal cord maintain the reciprocity of action between antagonist pools of flexor and extensor motor neurons. Unlike the sensory neurons of the stretch reflex, however, flexion-reflex afferents make only indirect connections to motor neurons, via at least one layer of intervening interneurons. The obvious function of the flexion reflex is protective; the offended limb is flexed, lifted, and withdrawn from a painful and potentially damaging stimulus. The reflex circuit is relatively short, local, and rapid. Of course, flexion-reflex afferents also connect to other interneurons that ascend the spinal column to the brain, so you become aware of the painful stimulus. This slower process occurs while the reflex flexion is taking place, so in most cases the foot is lifted (or the hand is withdrawn from the hot stove) before you are aware of the stimulus triggering the withdrawal. Note that many receptors other than pain receptors can trigger flexion reflexes, and that 4 the main function of the flexion-reflex afferents is to provide proprioceptive and cutaneous information to the brain and spinal cord, not just to elicit flexion reflexes. If you stepped on a tack with your left foot while your right foot was lifted off the ground, it would be a good idea to extend your right foot while flexing your left foot. In fact, one component of the flexion reflex ensures this. As Figure 19.2 shows, flexion- reflex afferents synapse onto interneurons that cross the midline of the spinal cord and indirectly excite extensor motor neurons of the contralateral (“opposite side”) leg. Thus the right leg is extended (by exciting extensor motor neurons and inhibiting flexor motor neurons) while the stimulated left leg is flexed (by exciting flexor motor neurons and inhibiting extensor motor neurons). The reflex extension of the contralateral leg has been given a separate name— the crossed extension reflex—but functionally it is an integral part of the flexion reflex, a product of the synaptic connections “wired in” to the spinal cord. This example illustrates that reflexes do not operate in a vacuum, influencing only a single antagonist pair of muscles. Instead, reflexes may have diverse and widespread effects, and they must interact with all other synaptic influences on motor neurons. The stretch reflex circuit illustrates the principle of divergence of central neural connections: Each presynaptic neuron usually contacts many postsynaptic neurons. The converse principle, convergence, also occurs, because each postsynaptic neuron is contacted by many presynaptic neurons. For example, each of the motor neurons receives input from about 10,000 synapses, representing many 1a sensory neurons and many more excitatory and inhibitory interneurons. Thus the cartoon view of the circuit for a stretch reflex in Figure 19.1 is a great oversimplification. To illustrate how spinal reflex circuits, provide sensory feedback, let’s consider how the stretch reflex compensates for a resistance or load during the execution of a centrally generated, voluntary movement. To emphasize that similar neural circuits control arms 5 and legs, we will use arm movements for this illustration. Suppose you decide to pick up a pamphlet from a table (Figure 19.4). Because this is a voluntary movement, the CNS must program the activation of motor neurons, rather than sensory input initiating the movement. Essentially the CNS estimates the amount of force necessary to pick up the pamphlet and sends a command to the motor neurons to generate that force. At the same time, the stretch reflex mediates load compensation, augmenting the contraction if there is extra weight or resistance added to the intended movement. The central command for a voluntary movement excites both alpha and gamma motor neurons, a process termed alpha-gamma coactivation. This coactivation has two functions. First, it ensures that the ongoing sensitivity of the muscle spindle is maintained during muscle shortening. In the absence of coactivation, a contraction that shortens the muscle would slacken the intrafusal muscle fiber and unload the muscle spindle, decreasing its sensitivity. Coactivation prevents this decrease. Second, coactivation allows the spindle to determine whether the muscle shortens during the intended movement. Suppose that the pamphlet is not very heavy, and the CNS has correctly estimated its weight. (We will call this the no-load condition, although it is more accurately described as light-load.) As shown in Figure 19.4a, the coactivation of and motor neurons activates contraction of both the intrafusal fibers and the extrafusal fibers of the working muscle. In the absence of a substantial load, the extrafusal muscle fibers shorten to flex the arm, allowing the intrafusal fiber associated with a muscle spindle to shorten as it contracts. The shortening of the intrafusal fiber as the motor neurons activate it decreases its tension and lessens its activation of the stretch receptor. Because shortening unloads the muscle spindle, the 1a afferent neuron of the stretch receptor generates few if any action potentials. That is, if the muscle shortens when it is signaled to shorten, no follow-up is needed. Now suppose that the pamphlet is very heavy and the force estimate of the CNS is 6 insufficient (see Figure 19.4b). (We will call this the loaded condition.) Coactivation excites the and motor neurons as before, but now the extrafusal muscle fibers do not shorten, because of the unanticipated load. In the absence of shortening of the whole muscle, excitation of an intrafusal fiber by motor neurons will activate the stretch receptor, producing a train of action potentials. This stretch receptor activity constitutes an error signal, a measure of how much the muscle failed to shorten as commanded. As Figure 19.4 shows, the stretch receptor’s 1a axon makes excitatory synaptic contact with a motor neuron that innervates the working extrafusal muscle fibers. Activity in the stretch receptor neuron (the error signal) excites proportional activity in the motor neuron, generating additional tension in the working muscle to overcome the load. The neuron pair of the stretch receptor and motor neuron is a reflex circuit that functions as a load-compensating servo loop, detecting an error (failure to shorten) and counteracting it (“more force, please”) within a centrally commanded movement. Let’s begin our exploration of the control of rhythmic behavior by asking: How does a locust fly? As Figure 19.5a shows, the movement of a single wing of a flying locust can be viewed as a simple up-and-down oscillation, generated by a set of elevator (or levator) and depressor muscles. The electrical activity of these muscles can be recorded from a tethered locust flying in a wind stream. This activity consists of alternating bursts of muscle potentials—the depressors being activated when the wings are up, and the levators being activated when the wings are down. Because each muscle depolarization results from an action potential in a motor neuron to that muscle, it is clear that flight results from the generation in the CNS of alternating bursts of action potentials in levator and depressor motor neurons. This kind of pattern—alternating bursts of activity in motor neurons to antagonist muscles— 7 underlies most forms of rhythmic behavior. How are the motor neurons to antagonist muscles activated in alternation to produce a rhythmic movement such as that of a locust wing? Historically, two kinds of hypotheses have been advanced to explain the neural basis of rhythmic movements: peripheral control and central control. According to the hypothesis of peripheral control, each movement activates receptors that trigger the next movement in the sequence. The position of a locust wing is monitored by several proprioceptors (see Figure 19.5a): a single wing-hinge stretch receptor that generates a train of impulses when the wing is elevated, and several other receptors that are activated when the wing is depressed. Locust flight could (in principle) operate by peripheral control, by having sensory feedback from wing sensory receptors activate the motor neurons for the next movement (Figure 19.5b). Thus elevation of the wings would excite the wing-hinge stretch receptor, which would synaptically excite depressor motor neurons, thereby lowering the wing. The lowered wing would terminate excitation of the wing-hinge stretch receptor and would excite the depression-sensitive receptors, which would synaptically excite levator motor neurons, elevating the wing and completing the cycle. The peripheral- control hypothesis is also called the chained-reflex hypothesis because each movement is a reflex response to sensory feedback resulting from the last movement. According to the hypothesis of central control, locust flight is sustained by a central pattern generator (CPG)—a neural circuit in the CNS that can generate the sequential, patterned activation of motor neurons to antagonistic muscles that underlies a behavior pattern, without requiring sensory feedback to trigger the next movement. 8 Thus, in central control of locust flight, the basic pattern of alternation of activation of levator and depressor motor neurons would result from an intrinsic CPG rather than from a chained reflex (Figure 19.5c). The crustacean stomach consists of two chambers: an anterior cardiac chamber containing teeth that function as a gastric mill to grind and chew food, and a posterior pyloric chamber containing a sieve that serves to keep food particles from passing to the rest of the gut until the particles are small enough. Figure 19.7 shows a simplified neural circuit and rhythmic output of one rhythm of the stomatogastric ganglion: the pyloric rhythm that controls the straining of food particles by the pyloric filter. The pyloric circuit (see Figure 19.7b) acts as a hybrid oscillator, containing an oscillator neuron (AB) that serves as the pacemaker for the rhythm. The oscillator cell is tightly electrically coupled to two pyloric dilator (PD) neurons so that these three burst together, inhibiting the other neurons in the network (see Figure 19.7c). The oscillatory AB neuron and the coupled PD neurons burst first, inhibiting follower cells (LP, PY, and two others not shown). At the end of the AB/PD burst, the LP cell recovers from inhibition faster than the PY cells; therefore, the LP cell bursts next and prolongs PY inhibition. PY neurons then burst and inhibit LP, until the next AB burst starts a new cycle. The pyloric circuit thus has both cellular oscillator and network oscillator properties. The generation of the pyloric rhythm depends primarily on the AB cellular oscillatory neuron, but its triphasic cycle (AB/PD LP PY ...) and timing depend on the strength and time course of inhibitory synapses and on intrinsic currents of the follower cells. 9 The rhythms and circuits of the stomatogastric ganglion exemplify another feature that may be of general importance: They are profoundly subject to modulation. The stomatogastric ganglion receives about 100 axons of neurons from other parts of the nervous system, many of which can secrete neuromodulators that act diffusely in the small ganglion to alter its motor output. At least 15 modulators are present, including the amines serotonin, dopamine, octopamine, and histamine; the classical transmitters acetylcholine and GABA; and several peptides, including proctolin, FMRFamide-like, and cholecystokinin (CCK)-like peptides. The most common effect of a neuromodulator is to initiate and maintain rhythmic activity in a network. For example, adding serotonin, octopamine, or dopamine to a previously quiescent isolated stomatogastric ganglion induces a pyloric rhythm (Figure 19.8), although the rhythms induced by the three modulators differ in detail. In general, the stomatogastric ganglion requires permissive modulatory input from extrinsic neurons for the expression of its rhythms. Many of the central and sensory neurons that provide this modulatory input are well characterized. The neuromodulators of the stomatogastric ganglion act in two ways: They alter the intrinsic membrane properties of individual stomatogastric neurons, and they alter the strengths and dynamics of synaptic connections of the neurons. Modulatory effects on intrinsic neuronal currents can induce cellular oscillation (many stomatogastric neurons are conditional oscillators), excite or inhibit particular neurons, or alter other excitable properties. Moreover, modulators can make individual synapses more or less potent, changing the functional circuit connections, as well as cellular activities. It is at once exhilarating and sobering to realize that neuronal circuits such as those of the stomatogastric ganglion are not rigidly “hard-wired,” but rather are plastic and malleable—exhilarating because the ability of neuromodulation to free a circuit from the “tyranny of the wiring” may underlie adaptive plasticity of neural control of behavior, but 10 sobering because of the realization that a circuit diagram such as that in Figure 19.7b is descriptive of only one state of a dynamically shifting circuit. Neurons can even shift from one functional circuit to another, firing in “gastric time” or “pyloric time” under modulatory influence, and circuit elements can combine to form new patterns of output. Growing evidence suggests that these roles of neuromodulators are of widespread importance among CPGs. The principles of central pattern generation and the interaction of central and peripheral control of movement were first developed from invertebrate studies, principally with arthropods and molluscs. In this section we consider the degree to which these principles also apply to vertebrates. We can start with the question: How does a cat walk? For the moment, let’s consider the cat nervous system as composed simply of three compartments that can influence movement: brain, spinal cord, and sensory input (Figure 19.10). The immediate generators of walking movements in a cat are the spinal motor neurons that control the limb muscles. The spinal circuitry associated with these motor neurons was introduced earlier. The motor neurons receive direct or indirect synaptic input from three sources: (1) descending input from the brain, (2) sensory input from proprioceptors and other receptors in the periphery, and (3) local input from intrinsic spinal circuits. If the spinal motor neurons are to be activated in the correct spatio-temporal pattern to produce walking, what are the roles of these three compartments in generating this pattern? 11 These experiments show that the brain does not need to provide timing information for walking. Noradrenergic fibers descending from the brain in intact cats presumably command or enable the expression of the walking pattern by spinal circuits, but they are not necessary for timing the stepping cycle of a limb; certainly injected L-dopa does not provide timing information. In other experiments, cats with brain sections (at level 1 in Figure 19.11) can walk on a treadmill when given un-patterned electrical stimulation to a mesencephalic locomotor command region. With increasing strength of stimulation, the rate of locomotion increases and the gait changes to a trot and finally to a gallop. Thus the brain may initiate locomotion and modulate it subject to conditions, but the brain is not necessary for generating the locomotor pattern. Sensory feedback from the hindlimbs is also unnecessary for hindlimb stepping movements, as can be shown by experiments similar to those just described. Cats with or without spinal transection (at level 2 in Figure 19.11) can make normally alternating stepping sequences following hindlimb deafferentation by cuts of the dorsal roots that contain the sensory afferent axons. (For the spinally transected cats, walking is initiated with L-dopa or clonidine.) These experiments indicate that the cat spinal cord contains a CPG for walking movements. Similar experiments indicate that fish, salamanders, toads, and turtles also have spinal locomotor CPGs. Sensory feedback can still play important functional roles in locomotion of intact vertebrates. Spinal reflexes stabilize and modulate the effects of centrally patterned locomotor output, but spinal reflexes themselves may also be modulated by the CPG. For example, the effect of mechanical stimulation of the top of the foot of a walking cat depends on the position of the foot in the stepping cycle. If the foot is off the ground and swinging forward, it is lifted higher when stimulated (“exaggerated flexion”). If the foot is on the ground and bearing the cat’s weight, the same stimulation produces a more forceful extension. This reversal of a spinal reflex (which is clearly adaptive for stable walking) shows that the central events of the stepping cycle can strongly modulate reflex function. The experiments described in this section demonstrate that the mechanisms of control of 12 rhythmic locomotor movements are fundamentally similar in many invertebrates and vertebrates. Although the cellular aspects may vary (e.g., different network mechanisms of central pattern generation), the functional roles of central and reflex aspects of control appear to be similar in many cases. Until recently, the production and control of complex motor functions have been substantially attributed to brain structures such as the cerebral cortex, basal ganglia, and cerebellum. In such views, the spinal cord was assigned a subservient function in the production of movement, playing a largely passive role of relaying the commands dictated to it by the brain. Many recent studies (including the locomotion studies described previously) provide evidence that the spinal motor circuits are active participants in several aspects of the production of movement, contributing to functions that had been ascribed to “higher” brain regions. Moreover, the roles of various brain areas in motor control can be difficult to separate from sensory, motivational, and other aspects of brain function. Views on motor control are changing as a result of new data and interpretations. We begin our examination of the execution of a voluntary movement with the motor areas of the cerebral cortex (Figure 19.13). The primary motor cortex (or simply motor cortex) lies just anterior to the central sulcus, a prominent valley in the convoluted cortical surface of most mammals. 13 Early studies demonstrated that electrical stimulation of areas of the primary motor cortex elicited movements of particular parts of the body, with a point-to-point correspondence between the area stimulated and the movements produced. Thus the body regions are represented on the surface of the primary motor cortex by a somatotopic map (Figure 19.14; see also Figure 15.8). The motor cortical somatotopic map was long viewed as a detailed representa- tion of individual body parts (such as digits of the hand) or even of individual muscles, but recent studies support a rougher, more complex map of movement patterns, organized to promote coordina- tion among muscles and joints rather than to control single muscles. The cerebellum is a large, highly convoluted structure at the dorsal side of the hindbrain. It is present in all vertebrates. The cerebellum regulates movement indirectly, adjusting 14 the descending motor output of other brain areas. The cerebellum is clearly involved in the coordination of movement, as demonstrated by the effects of cerebellar lesions in various animals, including humans. Voluntary movements are still possible following cerebellar lesions, but they are clumsy and disordered, lacking the smooth and effortless precision of normal movements. Movements are accompanied by tremor, and patients with cerebellar injuries report that they have to concentrate on each part of a movement, joint by joint. The cerebellum supports the smooth and coordinated execution of complex movements, by evaluating motor commands and sensory feedback to provide error correction signals for motor control during a movement. The cerebellum contains two major parts: an outer cerebellar cortex and underlying deep cerebellar nuclei. The sole output of the cerebellar cortex is to the deep cerebellar nuclei. Three functional divisions of the cerebellar cortex receive inputs from and project (send outputs) to different parts of the brain via different deep cerebellar nuclei: the vestibulocerebellum (posterior, interacting with the vestibular system), the medial spinocerebellum (coordinating ongoing movement via its output directed toward the motor cortex and brainstem motor nuclei), and the lateral cerebrocerebellum (concerned with motor planning as well as with planning and sequencing of non-motor cognitive behavior). The basal ganglia are a set of nuclei (clusters of brain neurons) located in the forebrain and midbrain, under the cerebral hemispheres. The most important areas (in terms of motor control) are the caudate nucleus, the putamen, and the globus pallidus. The caudate nucleus and putamen are similar in origin and function, and together they are termed the neostriatum (or simply striatum). The caudate nucleus and putamen receive ex- citatory input from many parts of the cerebral cortex, both motor and association areas. The caudate nucleus and putamen send inhibitory neurons to the globus pallidus. The major output of the basal ganglia is inhibitory; neurons from the internal segment of the globus pallidus inhibit neurons in the thalamus that excite the cerebral motor cortex. 15 The cellular architecture and synaptic interactions of the cerebellar cortex are elegantly precise and are as well-known as those of any other area of the brain. As Figure 19.15a shows, the cerebellar cortex contains five types of neurons and two principal types of input fibers. The axons of Purkinje cells constitute the only output of the cerebellar cortex; these end in the deep cerebellar nuclei below the cortical surface. The major synaptic interactions of the cerebellar cortex are shown in Figure 19.15a. Climbing fibers make powerful excitatory 1:1 synaptic contacts with Purkinje cells. Mossy fibers, in contrast, provide divergent excitatory input to many granule cells. Axons of granule cells ascend to the surface layer of the cortex and branch in opposite directions to become the parallel fibers, which make excitatory synaptic contacts with the other types of cerebellar cortical cells. The roles of climbing-fiber and mossy-fiber input are not completely clear. Climbing fibers are thought to convey error signals (sensory feedback from errors in movements), whereas mossy fibers may convey broader information about the sensory context of a movement. The synaptic interaction of parallel fibers and Purkinje cells is especially prominent. The parallel fibers pass through the flattened, planar dendrites of Purkinje cells at right angles (Figure 19.15b). Each Purkinje cell receives excitatory synapses from about 100,000 parallel fibers (in addition to 1 climbing fiber). Thus the climbing-fiber and mossy-fiber inputs differ greatly in the degree of divergence and convergence of their synaptic effects. 16 The basal ganglia are important in selecting movements, suppressing competing or unwanted movements, and initiating the selected movement. Most of the neurons in the circuits of the basal ganglia are inhibitory, so these functions involve considerable inhibitory interaction. Figure 19.16 shows how the basic synaptic connections of the basal ganglia function to disinhibit movement. The output of the globus pallidus pars interna (GP ) inhibits movements. For the initiation of a movement, this tonic inhibition is lifted, by disinhibition. The striatum (caudate nucleus and putamen) receives excitatory input from the cerebral cortex, and striatal neurons inhibit neurons in the globus pallidus via two pathways. In the direct pathway there are two inhibitory synapses: Striatal neurons inhibit GP , aid GP neurons inhibit neurons of the thalamus. Thus activation of striatal neurons inhibits i GP ieurons and disinhibits the thalamus, thereby allowing a movement (see Figure 19.16). The indirect pathway, in contrast, involves a chain of three inhibitory neurons: Striatal neurons inhibit neurons of the globus pallidus pars externa (GP ), eP neurens inhibit GP neurons, and GP neurons inhibit the thalamus. i i This triple inhibition means that striatal activity via the indirect path will suppress other activity in the thalamus, strengthening the tonic suppression of other unwanted movements and preventing them from competing with the movement selected. Description of the figure: In the direct pathway through the basal ganglia, neurons from the cerebral cortex excite striatal neurons, which inhibit neurons in the glo- bus pallidus pars interna. These globus pallidus neurons inhibit neurons in the thalamus that promote 17 movement. Cerebral cortical activation of striatal neurons transiently inhibits the tonic inhibitory output of the globus pallidus, disinhibiting the thalamus and activating a movement Excitatory synapses are indicated with a plus sign (+) and inhibitory synapses with a minus sign (–). As Figure 19.17 shows, the planning and programming of a movement can be viewed as separate from the execution of the movement. We can suppose that the decision to move starts in the association cortex (cortex that is not linked to any particular sen- sory or motor system), because the readiness potentials recorded prior to a movement are not localized to a specific cerebral area. Two loop circuits from the association cortex are thought to be involved in preprogramming a movement: one loop goes through the basal ganglia (selection and initiation) and another through the lateral cerebrocerebellum (initial programming). Both loops feed back to the motor cortex via the ventrolateral nucleus of the thalamus. The motor cortex then generates the appropriate pattern of activity to initiate the movement. Information about the command is sent to the spinocerebel- lum, via several subcortical nuclei. This process, termed command monitoring, “informs” the cerebellum of the intended movement. The spinocerebellum also receives ascending information—both sensory information about joint position and muscle tension, and central information from spinal and brainstem motor centers. The spinocerebellum may integrate this feedback information about the state of lower motor centers (internal feedback) and about the periphery (external feedback) with the monitored cerebral command. The cerebellar output can then modify and correct the command on a continuous basis as the movement evolves, using an integral of all relevant information (command, motor state, and sensory feedback). This continuous correction is presumably faster and smoother than, say, a correction system based on sensory feedback alone. There remains considerable controversy over the roles of all these brain areas in the control of movement, and the areas are also implicated in other functions in addition to 18 motor control. Even if the preceding description of brain area interaction in the execution of a voluntary movement is correct, it begs other questions (such as, how is a decision to move actually made?). Nevertheless, the relations shown in Figure 19.17 illustrate how brain areas may interact in planning, coordinating, and commanding movements in mammals. In humans, two movement disorders result from neurodegenerative changes in basal ganglia function: Parkinson’s disease and Huntington’s disease (Huntington’s chorea). Parkinson’s disease is characterized by difficulty in initiating movements (akinesia), so a simple task such as climbing stairs or getting up from a chair becomes almost impossible to carry out. Akinesia is often accompanied by postural rigidity and by tremors in limbs at rest. Huntington’s chorea represents the converse of parkinsonism: Movements occur uncontrolledly and are difficult to stop. Both chorea (uncontrolled but coordinated jerky movements) and athetosis (slow writhing movements) are associated with damage to the striatum. Box Extension 19.2 discusses these neurodegenerative disorders and suggests how they shed light on the normal generation of voluntary movements in humans and other mammals. How can degeneration of neurons in the basal ganglia produce such opposite effects? First we need to examine the neuronal circuitry of the normal basal ganglia in more detail. There are two major circuits on the basal ganglia, the direct pathway and the indirect pathway. Figure 19.16 in the text shows how the direct pathway promotes the generation of a voluntary movement by disinhibition. 19 The indirect pathway, as we noted, opposes this action by adding another layer of inhibition. The figure shows the location and circuitry of the basal ganglia. Normal and abnormal circuitry of the basal ganglia (1) Location of the basal ganglia in a cross-section of the human brain. The basal ganglia include the caudate nucleus and putamen (orange in the diagram; together called the striatum), the external and internal segments of the globus pallidus (red-violet), the subthalamic nuclei (green), and in the midbrain (not shown), the substantia nigra. The caudate nucleus and putamen, although anatomically separated from each other by white matter of the internal capsule, are functionally similar. (2) Basic circuit in the normal brain. Excitatory input (green) to the putamen (and caudate nucleus) activates neurons of the direct and indirect pathways. Neurons in the direct pathway inhibit (red arrow) neurons in the internal segment of the globus pallidus (GPi), and the GPi neurons in turn inhibit neurons in the thalamus and suppress movement generation by the motor cortex. In contrast, putamen neurons in the indirect pathway inhibit neurons in the external segment of the globus pallidus (GPe), and the GPe neurons in turn inhibit GPi neurons. Thus the direct and indirect pathways have opposing, but balancing, effects on the ease of movement generation. Modulating input from dopaminergic neurons in the substantia nigra pars compacta (SNc) enhances the direct pathways and inhibits the indirect pathway. Other circuitry through the subthalamic nucleus (STN) is not considered here. (3) In Parkinson’s disease the dopaminergic neurons of the SNc degenerate, unbalancing their modulatory effects on the striatum (putamen and caudate nucleus). Direct pathway output of the striatum is weakened and indirect output is strengthened, leading to excessive inhibitory output of GPi, suppressing movement generation. (4) In Huntington’s disease, striatal inhibitory neurons of the indirect pathway degenerate, leading to insufficient inhibitory output of GPi and excessive movement generation. Both Parkinson’s and Huntington’s diseases involve neuronal degeneration and altered neurotransmitter activity in the basal ganglia. In Parkinson’s disease, dopaminergic neurons in the substantia nigra degenerate. The synaptic endings of these dopamine neurons are in the striatum, so degeneration of the neurons deprives the striatum of dopaminergic input, which normally enhances the direct pathway through the globus pallidus and suppresses the indirect pathway. Parkinsonian symptoms, then, reflect an increased inhibitory output, leading to an inability to select and change behavioral activities. Dopamine replacement therapy (through provision of the dopamine precursor L-dopa, which crosses the blood–brain barrier) alleviates many symptoms of Parkinson’s disease, at least for a while. Implants of dopaminergic tissue into the striatal area may provide longer-lasting relief from Parkinson’s symptoms. In Huntington’s disease, in contrast, extensive loss of neurons in the striatum decreases striatal inhibition of the indirect pathway. Chorea then results from a decreased inhibitory output of the basal ganglia, decreasing the ability to suppress unwanted movements. 1 Muscle Chapter 20 Muscle is a tissue that consists of specialized contractile cells. All animal phyla have two categories of muscle cells: striated and smooth (or unstriated). Striated muscle cells have alternating transverse light and dark bands, giving them a striped appearance. The pattern of bands reflects the organization of the contractile proteins myosin and actin into regularly repeating units called sarcomeres. Smooth (unstriated) muscle cells also possess actin and myosin, but these proteins are not organized into sarcomeres. In vertebrates, striated muscles make up skeletal (attached to bones) and cardiac (heart) muscles. Smooth muscles of vertebrates are found primarily in hollow or tubular organs such as the intestine, uterus, and blood vessels. Invertebrates also have striated and smooth muscles, but they are not always found in the same distribution as in vertebrates. In arthropods, for example, the skeletal (attached to the exoskeleton) and cardiac muscles are both striated, but so are muscles of the alimentary (digestive) tract. Vertebrate Skeletal Muscle Cells 2 Skeletal muscles (Figure 20.1) are composed of bundles of long, cylindrical muscle fibers, or muscle cells (the two terms are used interchangeably). Connective tissue surrounds individual muscle fibers, bundles of fibers, and the muscle itself. It holds fibers together, provides a matrix for nerve fibers and blood vessels to gain access to the muscle cells, and contributes elasticity to the whole muscle. In addition, connective tissue weaves itself into tendons, which attach the muscle to bones, thereby transmitting force generated by the muscle fibers to the skeleton. Whereas small muscles may contain only a few hundred muscle fibers, large limb muscles of mammals contain hundreds of thousands of fibers. Single muscle fibers can be as long as 0.3 m (1 ft). Single fibers are typically 10 to 100 micrometers ( m) in diameter, although some (such as those in certain Antarctic fishes) can reach several hundred micrometers in diameter. Skeletal muscle fibers are multinucleate (contain many nuclei) because they form developmentally by the fusion of individual uninucleate cells called myoblasts. A muscle fiber is surrounded by a cell membrane sometimes referred to as the sarcolemma. (The prefixes myo- and sarco- both denote “muscle.”) Each muscle fiber contains hundreds of parallel, cylindrical myofibrils (see Figure 20.1b,c and the opening scanning electron micrograph). The myofibrils are 1 or 2 m in diameter and as long as the muscle fiber. Each myofibril has regularly repeating, transverse bands. The major bands are the dark A bands and the lighter I bands. In the middle of each I band is a narrow, dense Z disc, or Z line. The portion of a myofibril between one Z disc and the next Z disc is called a sarcomere (see Figure 20.1d). Thus one myofibril consists of a longitudinal series of repeating sarcomeres. The Z discs of adjacent myofibrils are lined up in register with each other, so the pattern of alternating A bands and I bands appears continuous for all the myofibrils of a muscle fiber. This alignment of banding within a muscle fiber gives the fiber its striated appearance. The striations are visible by light microscopy. Higher-magnification electron micrographs show that the myofibrils contain two kinds of myofilaments. The thick filaments (see Figure 20.1f ) are composed primarily of the protein myosin and are confined to the A band of each sarcomere. A single thick filament consists of 200 to 400 myosin molecules. The thin filaments (see Figure 20.1e) are composed primarily of actin. A single thin filament consists of two chains of globular actin molecules wrapped around each other in a helix. Thin filaments are anchored to proteins in the Z discs. They extend from the Z discs partway into the A bands of each flanking sarcomere, where they interdigitate with thick filaments. The central region of the A band, which contains only thick filaments and appears lighter than the rest of the A band, is called the H zone. A narrow dense region called the M line bisects the H zone. 3 Cross sections of a myofibril show the relationship of thick and thin filaments in a sarcomere (Figure 20.2). A cross section through the I band shows only thin filaments. A section through the part of the A band in which the thick and thin filaments overlap shows each thick filament surrounded by six thin filaments. A section through the H zone shows only thick filaments. A longitudinal view of a cylin- drical myofibril shows that it consists of a series of sarcomeres. Cross sections illustrate the regions of overlap of the thick and thin myofila- ments. Both the M line and the Z disc contain accessory proteins that anchor the thick and thin filaments. Top: This enlarged segment of the scanning electron micrograph in the chapter opener illustrates series of sarcomeres in six myofibrils. When a muscle fiber contracts, the thick and thin filaments do not shorten but instead slide by one another. Investigators in the 1950s noted that the A band, which is the length of the thick filaments, does not shorten during contraction. Furthermore, the distance between the margins of the two H zones on either side of a Z disc (the length of the thin 4 filaments) stays the same, no matter what the length of the adjacent sarcomeres. When a sarcomere changes length, the regions that also change length are the H zone and the I band. Two independent teams—A. F. Huxley and R. Niedergerke and H. E. Huxley and J. Hanson—made these observations in 1954 and formulated the sliding-filament theory of muscle contraction, which has since been amply confirmed. It states that the force of contraction is generated by the cross-bridges of the thick filaments attaching to the thin filaments and actively pulling them toward the center of the sarcomere Thick and thin filaments are polarized polymers of individual protein molecules Description of the figure: (a) About 200 to 400 myosin molecules polymerize to form a polarized thick filament. (b) Each myosin molecule contains two heavy chains of amino acids. The tail of the molecule consists of the two chains coiled around each other. The amino-terminal end of each heavy chain forms one of the heads. The head region has a surface for binding actin and a different site for binding and hydrolyzing ATP (ATPase activity). A link (or hinge) region of the heavy chain connects the head to the tail. In addition, the myosin molecule includes two smaller light chains associated with each head. Thus, each complete myosin molecule contains six polypeptide chains: two heavy and four light. The molecular composition of the heavy and light chains varies in different types of muscles. The different myosin isoforms of heavy chains and light chains confer variations of functional properties, such as the rate at which the myo- sin ATPase hydrolyzes ATP. The myosin molecules that produce contractile force in smooth and striated muscles are classified as myosin II. They are part of the myosin superfamily that consists of at least 18 different classes of myosins found in protozoans, fungi, plants, and animals. Individual myosin molecules are large proteins of about 500 kilo- daltons (kDa), each consisting of two globular heads joined to a long rod, or tail. The heads are the cross- bridges, and the tail con- tributes to the backbone of the thick filament (Figure 20.4). During polymerization the myosin molecules orient themselves with their tails pointing toward the center of the thick filament and their heads toward the ends. As a result, the two halves of the thick filament become mirror images of each other with a short bare zone of only tails in the middle of the filament. The cross-bridges on either side of the 5 bare zone point in opposite directions. Each actin molecule is a globular protein (42 kDa) called G- actin. G-actin monomers form chains of F-actin (filamentous actin). The two chains of F-actin wind around each other in a helix (see Figure 20.1e). Like the myosin molecules in thick filaments, G-actin molecules in thin filaments are arranged so that those on one side of the Z disc have one orientation, and those on the other side have the opposite orientation. The consequence of the polarized organization of the thick and thin filaments is that the cross-bridges in contact with the thin filament can act like oars to pull the thin filaments toward the center of the sarcomere. Myosin heads cyclically attach to actin molecules and then swivel to pull on the actin filament. Each myosin head has two binding sites: one for actin and the other for ATP. The binding site for ATP is an ATPase with enzymatic activity that splits inorganic phos- phate from the ATP molecule and captures the released energy. The energy is used to power cross-bridge action. The cycle of molecular interactions underlying contraction is shown in Figure 20.5. In step 1 the myosin head is bound to actin but is not binding ATP. This is the rigor conformation, as in rigor mortis, in which muscles of a dead person (or other animal) are rigidly fixed in place because of the absence of ATP in death. ATP is required for myosin to unbind from actin (step 2). In life, the rigor stage of each cross-bridge cycle is brief because the globular myosin head readily binds ATP, which causes the myosin head to detach from actin. It is important to understand that the detach- ment of myosin from actin requires the binding of ATP to change the conformation of myosin’s actin-binding site, but it does not require the energy derived from the ATP. 6 Once released from actin, the myosin head hydrolyzes the ATP to ADP and inorganic phosphate (P i (step 3). A change in angle of the myosin head (termed cocking) accompanies hydrolysis, but the ADP and P remiin attached to the head. The energy released by hydrolysis of ATP is stored in the myosin–ADP–P complex. The i complex then binds actin (step 4), forming an actin–myosin–ADP–P complei. Initially actin binding is weak, but it triggers P release, tighter binding, and the power i stroke (step 5). The myosin head swivels, pulling the attached actin toward the middle of the myosin filament. At the end of the power stroke, the ADP is released and the myosin remains tightly bound to the actin (step 6). A new molecule of ATP then binds to the myosin head, triggering its release from actin. In a resting muscle, each myosin head has detached from actin, hy- drolyzed the ATP, and stored the energy obtained from hydrolysis. It is “primed” for another cycle. However, the regulatory proteins tropomyosin (TM) and troponin (TN) prevent contraction by inhibiting most of the myosin heads from binding to actin. TM is a protein dimer of two polypeptides that form an -helical coiled-coil, which lies along the groove between the two actin chains of the thin filament (Figure 20.6). A single TM molecule extends the length of seven globular actin molecules. Each TM molecule is associated with one TN molecule. TN is a golf club–shaped complex of three subunits. The “handle” is troponin T (TN-T), which binds to tropomyosin. The “club” includes 2+ troponin I (TN-I), which binds to actin, and troponin C (TN-C), which binds Ca ions. In the resting state (see Figure 20.6a), the TM molecule lies over the myosin-binding 7 sites of the adjacent actin molecules and prevents myosin cross-bridges from binding to actin. For contraction to occur, TM’s inhibition of cross-bridge binding is counteracted by the binding of Ca2+ to TN. The key physiological regulator of muscle contraction is calcium. 2+ When Ca ions bind to TN-C, they trigger conformational changes in TN, TM, and probably actin, which permit myosin cross-bridges to interact with actin (see Figure 20.6b). The changes that occur when Ca2+ binds to TN appear to involve both removal of TM’s steric blocking of the myosin-binding sites on actin and also more subtle allosteric interactions among the proteins. Once interaction between actin and myosin is possible, the primed myosin cross- bridges are permitted to go through cross-bridge cycles until the Cis removed. The muscle 2+ will therefore contract only when Ca ions are available to bind TN. In relaxed skeletal muscle fibers, the intracellular concentration of calcium is extremely –7 low: less than 1 10 M, which is below the concentration that will induce (by mass action) calcium association with troponin. 8 ATP is the immediate source of energy for powering muscle contraction ATP performs at least three functions in the contraction–relaxation cycle: 1. ATP binding to the cross-bridge (but not hydrolysis) is necessary for detachment of myosin from actin. 2. Hydrolysis of ATP activates (cocks) the myosin cross-bridge in preparation for binding to actin and undergoing a power stroke. 2+ 3. Energy from the hydrolysis of ATP drives the calcium pump that transports Ca ions into the sarcoplasmic reticulum. However, muscle contains only enough ATP (2–4 mM) to sustain contraction for a few seconds. Thus, nearly all forms of muscular work depend on regeneration of ATP while the muscle is working. The rate of muscular work strictly depends on the rate at which ATP is provided to the contractile apparatus. In broad outline, vertebrate muscle fibers possess three biochemical mechanisms that produce ATP: 1. Use of the phosphagen creatine phosphate. Phosphagens temporarily store high- energy phosphate bonds. The high- energy phosphate of creatine phosphate can be donated to ADP to produce ATP. Creatine phosphate is produced in resting muscle from creatine and ATP. The formation of ATP from creatine phosphate is driven by simple mass action. Whereas creatine phosphate is the phosphagen found in muscles of vertebrates, it and other phosphagens, such as arginine phosphate, are found in invertebrates. 2. Anaerobic glycolysis. This form of catabolism requires no oxygen. It must have glucose or glycogen as fuel. In addition to ATP, it produces lactic acid, which in vertebrates is always retained in the body and disposed of metabolically. 3. Aerobic catabolism. This form of catabolism requires oxygen and can use all three major classes of foodstuff as fuel. It produces ATP principally by oxidative phosphorylation. Its other major products are CO2and H O2 Figure 20.13 summarizes major elements in the production and use of ATP in a vertebrate muscle fiber. As emphasized in Chapter 8, the three mechanisms of ATP production differ greatly in how fast they can make ATP when operating at peak output, how much ATP they can make, and how quickly they can accelerate their rate of ATP production. 9 In this transverse section, small-diameter slow oxidative fibers (red) stain dark because of their abundant mito- chondria, and they are surrounded by many capillaries (black). Large- diameter fast glycolytic fibers (yellow) have fewer mitochondria and few immediately adjacent capillaries. Fast oxidative glycolytic fibers (orange) are intermediate in diameter and in abundance of capillaries. 10 A vertebrate skeletal muscle is innervated typically by about 100 to 1000 motor neurons. The axon of each motor neuron typically branches to innervate multiple muscle fibers, and each muscle fiber receives synaptic input from only one motor neuron. A motor neuron and all the muscle fibers it innervates are collectively termed a motor unit (Figure 20.15). When the motor neuron generates an action potential, all of the muscle fibers in the motor unit generate action potentials and contract to produce a twitch. Trains of action potentials of increasing frequencies can produce summation of twitches up to fused tetanic contraction. Thus, as in whole muscles, the amount of tension produced by a single motor unit can be varied by varying the frequency of action potentials generated by the motor neuron. Although the amount of tetanic tension varies in different animals, in many vertebrate muscles it is only two to five times the twitch tension. A more dramatic effect on the amount of tension developed by a whole muscle can be accomplished by varying the number of active motor units. Increasing the number of active motor units is called recruitment of motor units. Recruitment requires stimulating increasing numbers of motor neurons that innervate the muscle. For example, the tension in a muscle innervated by 100 motor neurons could be graded in 100 steps by recruitment. The amount of tension developed by the whole muscle increases as more motor units are activated (recruited). Recruitment is the dominant means used to control the amount of tension produced in vertebrate twitch muscles. Varying the number of active motor units, as well as the timing of their activation, ensures precise and smooth movements. The elastic properties of the muscle also contribute to the smoothness of movement. 11 Whereas each fiber of a twitch muscle has a single synaptic contact near the middle of the fiber, each muscle fiber of a tonic muscle receives many branches of a motor neuron, so it has many synaptic contacts distributed over its length. This pattern, shown in Figure 20.16a, is termed multiterminal innervation. An action potential generated by a motor neuron produces an excitatory postsynaptic potential (EPSP) at each of the distributed junctions. The muscle fiber has little or no ability to generate action potentials. Each depolarizing EPSP spreads passively over a region of membrane and down the t-tubules in that area. Contraction occurs by excitation–contraction coupling. Because an EPSP is produced at each of the many terminals along the entire length of the fiber, the contractile elements along the entire fiber are activated. The amount of tension generated depends directly on the amount of depolarization produced by the EPSPs. Although the fibers of arthropod skeletal muscles share many features of vertebrate skeletal muscle, including the organization of thick and thin filaments into sarcomeres and excitation–contraction coupling by way of t-tubules and SR, they show interesting differences in their patterns of innervation. A typical arthropod whole muscle is innervated by one to ten motor neurons, in contrast to the hundreds or thousands of motor neurons that innervate a whole vertebrate muscle. Most individual arthropod muscle fibers are innervated by more than one motor neuron, a pattern termed poly-neuronal innervation (Figure 20.16b). 12 One useful classification scheme differentiates vertebrate smooth muscle into two main types: single-unit and multiunit smooth muscles (Figure 20.17). In single-unit smooth muscle, the muscle cells are tightly electrically coupled by numerous gap junctions (see Figure 20.17a). Because of this coupling, groups of muscle cells are de- polarized and contract together, functioning as a single unit. The smooth muscles of the gastrointestinal tract and small- diameter blood vessels are examples of the single-unit type. Single-unit smooth muscle is often spontaneously active, with electrical activity propagating from cell to cell via the gap junctions. This type of muscle can also be activated by stretch. Neural and hormonal controls may modulate the endogenous activity to varying degrees. Multiunit smooth muscles have few if any gap junctions, so the muscle cells function as independent units (see Figure 20.17b). They are innervated by aut
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