The Neural Control of Contraction in a Fast Insect Muscle ROBERT K. JOSEPHSON, DARRELL R. STOKESZ AND VICTOR C H E N 3 School of Biological Sciences, U n i v e r s i t y of C a l i j o m i a , I r v i n e , California 92664

ABSTRACT The wing muscles used in singing by the katydid, Neoconocephalus robustus, are extraordinarily fast. A t 3 5 " C , the animal's thoracic temperature during singing, an isometric twitch lasts only five to eight msec (onset to 50% relaxation) and the fusion frequency of these muscles is greater than 400 Hz. Stimulating the motornerve to a singing muscle initiates a short (2.5 msec at 3 5 " C ) , sometimes overshooting depolarization of the muscle fibers. Despite their spike-like appearance, the electrical responses are largely synaptic potentials. The muscle membrane appears to be capable of only weak, electrically-excitable, depolarizing electrogenesis. The short synaptic potentials result in part from rapidly-developing delayed rectification, in part from a low resting membrane resistance (Rm = 162 RcmZ) and a concomitantly short membrane time constant (about 1.5 msec).

Adult male katydids (Orthoptera, Tettigoniidae) stridulate by rubbing a sclerotized ridge on the medial edge of one forewing against a set of teeth on the underside of the other forewing. In some cone-headed katydids (Subfamily Copiphorinae) the wing stroke frequency during stridulation can be extraordinarily high, 160 Hz in Euconocephalus n a s u t u s (Josephson, '73) and up to 212 Hz in Neoconocephalus robustus (Josephson and Halverson, '71). Unlike some high frequency muscles of insects, the singing muscles of katydids are neurogenic, there is an electrical potential from the muscle preceding each contraction. The ability of the singing muscle to operate at high frequency depends on its ability to contract and relax rapidly following each arriving nerve impulse and not to intrinsic oscillatory properties of the muscle. The high frequency performance during singing is achieved in part by operating the muscles at elevated temperature. In both E . nasutus and N . robustus the thoracic temperature during singing is about 35 C when the environmental temperature is 25 C (Heath and Josephson, '70; Josephson, '73). But even allowing for the elevated temperature, activation processes in a singing muscle are extremely fast. At 35" C the contraction time for an isometric twitch of a singing muscle from E. nasutus is only 4.6 O

O

J . ExP. ZOOL., 193: 281-300

msec and the time from twitch onset to 50% relaxation is 7.7 msec. For comparison, twitch durations (onset to 50% relaxation) for some fast mammalian and bird muscles at a similar temperature are: rat inferior rectus, 9.2 msec (Close and Luff, '74); cat inferior oblique, 12-15 msec (Bach-y-Rita and Ito, '66); bat stapedius, 11 msec (estimated from attenuation of cochlear microphonics, Henson, '65); zebra finch flight muscle, 16 msec (Aulie and Enger, '69); hummingbird pectoralis major, 16 msec (measured from figure in Hagiwara et al., '68). The activation of striated muscle involves a number of steps; including depolarization of the surface membrane, inward spread of excitation along the transverse tubular system, release of calcium from the sarcoplasmic reticulum, diffusion of calcium into the myofibrils, and binding of calcium to the myofilaments (Ebashi and Endo, '68; Huxley, '71; Weber and Murray, '73). The diffusion of calcium from the sarcoplasmic reticulum to the myofilaments (and back during relaxation) is possibly one of the 1 This work was supported by NIH Grant NS-10336. The authors wish to thank Drs. H. Y. Elder and F. Lang for their comments o n a n earlier version of this paper. Present address: Department of Biology, Emory University, Atlanta, Georgia 30322. 3 Present address: Department of Biology, Case Weste m Reserve University, Cleveland, Ohio 44106.

28 1

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ROBERT K. JOSEPHSON, DARRELL R . STOKES AND VICTOR C H E N

rate limiting steps. Fast muscles are characterized by an extensive sarcoplasmic reticulum ramifyin g throughout the muscle . This feature minimizes the distance and hence diffusion time between sarcoplasmic reticulum and myofilaments. In the singing muscles of katydids the sarcoplasmic reticulum is unusually well developed, and comprises 1%25% of themuscle volume (Elder, '71; Stokes et al., '75). At the very short contraction times encountered in katydids, the depolarization of the surface membrane is also a potential rate limiting step. Arthropod muscles are characterized by multitermjnal innervation and stimulation of a n excitatory motorneurone leads to distributed excita tory po st-synap tic potentials (EP SP's) in the muscle fibers which the motorneurone innervates. The EPSP's may give rise to elec trically-excited membrane responses which are often graded in amplitude (Usherwood, '69; Atwood, '72). In some crustacean muscles the synaptic potentials can trigger all-or-nothing muscle action potentials (e.g., Fatt and Katz, '53, Atwood, '65, Ozeki, '69; Hoyle and Burrows, '73). Electrically-excited, all-or-nothing action potentials have not been unequivocally demonstrated in insect muscle (Usherwood, '69). The electrical responses of the muscle membrane following nerve stimulation can be reasonably short, 3.5-4 msec (1617°C) in lobster sound-producing muscle (Mendelson, '69) and about 4 msec (1% 20°C) in coxal muscles of the cockroach (Becht et al., '60). These are unusually brief responses; in most arthropod muscles which have been investigated the electrical response to nerve stimulation lasts 5-10 msec or more (e.g., Werman et al., '61, Pamas and Atwood, '66; Hoyle and Burrows, '73). Even %4 msec is a n appreciable fraction of the extremely short twitches of katydid singing muscles. The rapid activation and inactivation of the singing muscle would seem to require very brief membrane depolanzation as a n initiating step in the contraction sequence. The following investigation was begun to determine in what ways, if any, the membrane properties of the singing muscle are specialized to promote rapid muscle activation and inactivat ion. MATERIALS AND METHODS

The animals used were adult, male Neo-

conocephalus robustus collected from salt marshes in the vicinity of Woods Hole. Most experiments were done with the dorsal longitudinal muscle (DLM). This broad, flat muscle was chosen because it is the thinnest of the major flight and singing muscles. The thinness of the DLM allows access to all of its fibers with microelectrodes and, in addition, i t should facilitate exchange of respiratory gases between the muscle and its environment. In the usual preparation the legs and wings were removed. The gut was exposed in the neck, ligated, and cut anterior to the ligation. The portion of the gut in the thorax and abdomen was then removed through a slit in the abdominal wall. The hole in the neck was filled with a saline-soaked wick which later served as a reference point in electrical recording. The saline which was used was Usherwoods ('68) modification of Hoyle's locust saline. The animal was fastened to a dissecting dish using staples which bracketed but did not penetrate the body. The DLM and its motornerve were exposed by removing the lateral body wall and the overlying dorsc-ventral muscles, care being taken to leave the trachea to the muscle as intact as possible. The muscle was activated by stimulating its motornerve with a suction electrode. Muscle tension was recorded with a n RCA 5734 mechanc-electric transducer. The second phragma, the posterior insertion of the muscle, was stabilized by placing the horizontal bar of a staple against the anterior face of the phragma. A short steel pin attached to the anode of the transducer was placed against the posterior face of the first phragma, the anterior insertion of the mesothoracic DLM. The pin was sufficiently stiff that contraction was essentially isometric. The transducer was mounted in a manipulator so that muscle length and resting tension could be varied. It is likely that removal of the support offered by the lateral body wall allows the muscle in dissected preparations to shorten somewhat from its usual in vivo length. The performance of katydid muscles does vary with muscle length; the twitch tension increases and the twitch time course becomes longer with increasing muscle length to lengths 40% or more greater than slack length (Josephson, '73). In our measurements the muscle was

CONTRACTION I N A FAST I N S E C T MUSCLE

stretched to a length 10% greater than slack length. At this length, which is close to or perhaps a bit longer than the normal in vivo length, twitch tension is less than maximal but the twitch time course is nearly minimal. The force produced by the contracting muscle is distributed along the phragma, while it -was measured from one point near the middle of the insertion. Different parts of the muscle pull against the transducer through lever arms of different lengths. The effective fulcrum of the phragma in these preparations is uncertain and it is therefore not possible to evaluate possible effects of varying lever arm length. Different parts of the muscle may thus vary in their ability to transmit force to the transducer because of the position of their insertion as well as because of possible intrinsic differences in contractile strength. Because of this, no attempt has been made to calculate the force produced by the muscle or its parts in terms of tension per unit cross-sectional area. The muscle temperature was varied by warming the whole animal with a heat lamp or with a microscope illuminator placed close to the animal. The temperature of the blood surrounding the muscle was measured with a thermocouple or a small thermistor. The temperature probes used were approximately 0.3 mm in diameter. Membrane potentials from muscle fibers were recorded with glass capillary microelectrodes filled with 3 M KCl and with resistances ranging from 10 to 30 MR. The muscle fibers are small, 20-35 pm in minimum diameter (Stokes et d.,'75) and muscle movement made stable electrode penetration a bit difficult. In addition to the rapid twitches following stimulation, the muscle was displaced laterally by several hundred pm during breathing movements of abdominal and remaining thoracic musculature. Successful results were obtained with three different electrode arrangements: (1) floating single electrodes. The microelectrode was suspended from a length of polyethylene tubing whose central portion had been heated over a small flame and pulled out to a fine (ca. 100 pm diameter), hollow filament (Benoit, '62: fig. 1). The polyethylene tubing was filled with 3 M KC1. The stem of a microelectrode was

283

scored and broken off close to the shoulder. The shortened electrode was inserted into one end of the polyethylene tubing. The other end of the polyethylene tubing fit over a piece of glass capillary in a leucite electrode chuck which, in turn, was held in a manipulator. In this way the electrode was attached to the manipulator through a thin, very flexible, salt bridge. In use the electrode was positioned over the muscle and lowered until i t penetrated a fiber. In most cases the electrode remained in the fiber during both the rapid accelerations of evoked twitches and slow breathing movements. Potentials were often recorded which are thought to be movement artifacts because they coincided with muscle contraction (fig. 4), but these were generally small enough that they posed no major problem. The length and diameter, and hence the flexibility, of the polyethylene tubing could be easily varied in the pulling stage. It might be pointed out that using a flexible salt bridge avoids junctional potentials which can occur when microelectrodes are floated directly from fine metal wires (2) double-barreled floating electrodes. Short, double-barreled electrodes were made in a puller similar to that described by Mendelson ('67). One of the barrels (the recording channel) was suspended from a drawn polyethylene segment. A fine silver wire (25 pm diameter), connected to a current source, was inserted into the other barrel which was used to inject current into the cell. The injected current was monitored with an operational amplifier as a current to voltage converter in the ground return from the preparation (Moore, '63) ( 3 ) independent, rigidly-mounted electrodes. Some problems with inter-electrode coupling were encountered when using double-barreled electrodes (see below). To minimize electrode coupling some fibers were penetrated with two independent electrodes, one for passing current and the other for recording potential. Current was again monitored with an operational amplifier. The electrodes were placed close together, usually less than 100 pm apart. Precise electrode positioning required rigidly-mounted electrodes and muscle movement was particularly troublesome here. In these experiments the ganglion of the mesothorax was removed to paralyze the remaining intact

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ROBERT K. JOSEPHSON, DARRELL R. STOKES AND VICTOR CHEN

Plastic Tubing

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Microelect rode

Fig. 1 The method used to flexibly mount microelectrodes. The figure is not to scale, in use the tubing section was considerably longer and thinner than is shown.

electrodes in its dorsal origin and tension was measured isometrically with the muscle approximately 10 % longer than its slack length. The muscle temperature was 25°C and 3 5 " C , the latter being approximately RESULTS the thoracic temperature during singing in A n interspecies calibration both species. Some twitch parameters from homoloFor reasons given above, the muscle examined most closely in N. robustus was the gous muscles in N. robustus and E . nasutus mesothoracic DLM. But previous mechan- are summarized in table 1. The data is ical measurements from katydid muscles based on only three muscles from N. robuswere from another muscle in a different tus but measurements were made from 10 species; the first tergocoxal muscle of Euco- replicate twitches from each muscle at the nocephalus nasutus (Josephson, '73). In two temperatures. While muscle performorder to better compare the present results ance in the two animals is generally simwith the earlier ones, some tension mea- ilar, it is clear that the muscles of N. robussurements were made from the first tergo- tus are somewhat faster than those of E . coxal muscle of N. robustus. The techniques nasutus. Probably related to this, the twitch used were the same as with E. nasutus. tension in N. robustus is less than that of The muscle was stimulated directly through E . nasutus. This is similar to results from

muscles of the segment and the DLM was further immobilized by bracketing it with staples which pressed firmly against the muscle surface.

285

CONTRACTION IN A FAST INSECT MUSCLE TABLE 1

Comparison of isometric t w i t c h contractions of mesothorucic f i r s t tergocoxal m u s c l e s f r o m N. robustus a n d f r o m E. nasutus, m e a n ( r a n g e ) N . robustus n = 3 2 5’

Rise time, msec Onset to 50% relaxation, msec Onset to 90% relaxation, msec Twitch tension, gm c m - *

350

6.6

(6.2-6.8) 11.1 (10,&11.8) 20.0 (17.2-22.1) 26

(14-39)

homologous singing and non-singing muscles in E. nasutus in which there is an inverse relation between contraction speed and tension development (Josephson, ’73). The faster twitches of N. robustus are consistent with the faster wing stroke frequency during singing of this species; the mean wing frequency during singing of N. robustus is 187 Hz (Josephson and Halverson, ’71) while that measured for E. nasutus was 157 Hz. The faster performance of N. robustus muscles extends to their responses to repetitive stimulation. At 35°C an E. nasutus singing muscle is able to relax by 53% between tension peaks when stimulated at 150 Hz, approximately the singing frequency (Josephson, ’73). With N. robustus muscles at the same temperature and frequency, the tension in the troughs is only 21% of that at the peak (n = 3, range = 1525%). At 200 Hz, approximately the singing frequency of N. robustus, the muscle relaxes about 53% between tension peaks (n = 3, range = 40-67%). A t 3 5 ° C the N. robustus muscle follows repetitive stimulation to at least 500 Hz and tension fusion is obviously incomplete at 400 Hz (fig. 2).

Electrical and mechanical activity of the DLM Stimulating the nerve to the DLM with single shocks of gradually increasing intensity produces four discrete tension increments, indicating that the muscle is innervated by four, twitch-type motorneurones (fig. 3A). Confirming this, cross sections of the nerve to the DLM show four major axon profiles (fig. 3B). Three of these motorneurones produce twitches with similar time

E . nasutu s n = 6

3.1 (2.8-3.3) 5.1 (4.3-5.5) 8.5 (6.7-10.5) 40

(15-69)

250

350

7.1 (6.S8.0)

(4.0-5.9)

4.6

12.2

7.7

(10.8-15.2)

(6.9-10.2)

20.0

12.2

(16.1-29.3) 74 (42-94)

(10.8-16.5) 80

(42-1 07)

course, the fourth (henceforth called the slower axon) initiates a distinctly slower twitch. In some preparations the slower axon had the lowest threshold, in other preparations it had the second or rarely the third lowest threshold. It should be pointed out that the slower axon is not a “slow axon” as this term is usually applied to insect preparations (e.g., Usherwood, ’62). Motoraxons described as “slow axons” typically give rise to small, facilitating electrical responses of the muscle and very slow contraction. The slower axon of the katydid DLM initiates large electrical responses of the muscle fibers which it innervates (Stokes et al., ’75) and the twitches which it evokes are comparable in speed to the fast twitches of, for example, locust flight muscle (Neville and Weis-Fogh, ’63). The responses to the slower axon of N. robustus are slow only in comparison with the extraordinarily fast responses initiated by the other axons. The slower axon to the DLM innervates only the most dorsal portion of this muscle (Stokes et al., ’75). Electrical recordings considered in this paper are from ventral and medial portions of the DLM and so represent responses of only the faster axons. The resting potential of muscle fibers was typically between - 30 and - 50 mV. This seems a bit low, suggesting that the fibers may have been damaged by electrode insertion, but it is about the same as membrane potentials from muscles of grass-fed locusts when the muscles were bathed in locust hemolymph (Hoyle, ’54). In our study saline was added sparingly to the preparation and the muscle was bathed in a mixture of saline and hemolymph. If the hemo-

286

ROBERT K. JOSEPHSON. DARRELL R. STOKES AND VICTOR CHEN

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lymph had a high potassium level, reflecting the katydid's herbivorous diet, it might account for the seemingly low resting potentials. Stimulating the motornerve initiates a fast, sometimes overshooting electrical response in muscle fibers (fig. 4). Characteristics of these electrical responses and associated twitches are summarized in table 2. The total duration of the electrical response is about 4 msec at 25"C, about 2.5 msec a t 35°C. Evidence presented below suggests that, despite its short time course, the electrical response is largely a synaptic potential. Temporal characteristics of twitches from the DLM are generally similar to those of the first tergocoxal muscle. The heterogeneity of units in the DLM makes its response to repetitive stimulation more complex than that of the tergocoxal muscle. During repetitive stimulation a t moderately high frequency and with maximal stimuli, the mechanical responses initiated by the slower axon sum, and relaxation of the faster units is, at best, to the level of the rising tension created by the slower unit (fig. 5). The low threshold of the slow axon makes i t impossible to selectively activate all the fast units alone by varying the stimulus intensity. The best muscle performance in terms of fractional relaxation during repetitive stimulation occurs with stimuli which activate one or two fast axons and are below threshold for the slower axon. In our best preparations, muscles at 35°C relaxed by 50% or more between tension peaks when stimulated a t 200 Hz with stimuli of intermediate intensity. The maximum relaxation recorded at 35°C and 200 Hz was slightly more than 85% (fig. 6). Many preparations performed poorly to repetitive stimulation, relaxing only 10 or 20% between tension peaks a t 200 Hz and 35°C. Sometimes this poor relaxation was clearly due to asynchrony between individual motor units for the tension records contained multiple peaks for each stimulus; in other instances the reasons for the longer contractions were not apparent. The muscle performance generally deteriorated with time and only fresh preparations behaved like that of figures 5 and 6. The performance of singing muscles is easily degraded (Josephson, ' 7 3 ) and the extensive surgery needed to expose the DLM may have adversely affected its ability to

28 7

CONTRACTION IN A FAST INSECT MUSCLE

A

2 0 msec

20jm

Fig. 3 A . Mechanical responses of the DLM following nerve stimulation with a number of shocks of gradually increasing intensity, T = 25'C. The presence of four tension increments indicates that the muscle is innervated by four motorneurones. For three of the tension increments relaxation is essentially complete i n 15 msec. T h e tension step initiated by the fourth axon, here with the second lowest threshold, is considerably longer. B. Cross section of the motornerve showing four large axon profiles. The smaller profiles may be sensory axons.

i 10 msec Fig. 4 Electrical response of a DLM fiber (middle trace, zero potential indicated by upper line) a n d the associated twitch from t h e whole DLM (lower trace) following a single maximal shock to the motornerve, T = 25'C.

288

ROBERT K JOSEPHSON, DARRELL R STOKES AND VICTOR CHEN TABLE 2

Electnccil w i d m e c h a n t c u l chnractenstics of t w i t c h e s f r o m t h e DL m u s c l e of N robustus, metrn ( r a n g e ) . n zndictrtes t h e n u m b e r of preparations T h e rctnge given is t h a t of t h e m e a n s of t e n obseruutions f r o m e a c h preparutzon In the rlectrzccil pcirumeters e a c h observrrtzon wusf q o m n different m u s c l e fiber 25°C

35oc

Mechunicul response Rise time, msec ( n = 6) Onset to 50'% relaxation, msec ( n = 6) Onset to 90% relaxation, msec (n = 5) Tension, mg ( n = 6)

4.2 (3.7-4.8) 12.8 (8.517.0) 26.2 (14.1-38.3) 68.5 (52.3-84.8)

3.4 (2.44.3) 7.6 (6.3-9.0) 15.2 ( 9 . s 2 0 . 6 ) 78.2 (62.S93.5)

Electrical response Resting potential, mV ( n = 7) Response height, mV ( n = 7) Stimulus to response onset, msec ( n = 7) Rise time, msec ( n = 7) Onset to 50% return, msec ( n = 7) Onset to 90% return, msec ( n = 6 )

43.1 (39.446.7) 37.2 (23.9-50.5) 2.0 (1.7-2.4) 1.4 (1.1-1.6) 2.3 (1.7-2.9) 4.0 (2.7-5.3)

43.2 (37.548.9) 36.9 (24.749.1) 1.3 (O.Sl.8) 0.9 (0.7-1.0) 1 . 5 (1.1-1.9) 2.5 (1.4-3.6)

contract and relax rapidly. We therefore feel that the performance of our fastest preparations is most representative of the muscle capability in intact animals. The DLM often shows pronounced mechanical facilitation during repetitive stimulation (fig. 6). Surprisingly, facilitation of twitch height is not accompanied by facilitation of the electrical responses which, in fact, get smaller. During repetitive stimulation there is a small depolarizing shift in the potential baseline and diminution of electrical response amplitude. Thus late in a train of stimuli a smaller electrical response initiates a much larger twitch than at the onset of the train. The possibility that electrical responses somewhere in the muscle do facilitate during repetitive stimulation cannot be ruled out but we consider this unlikely. All the fibers which we impaled, including examples from dorsal, medial and ventral parts of the muscle, behaved like those of figure 6 with large electrical responses to the first shock and declining responses to subsequent stimuli. Electrical properties of muscle fibers measurements w ith doublebarreled electrodes The basic question considered in this and the following section is whether the recorded electrical responses are synaptic potentials, or spikes in an electrically-excitable membrane, or a combination of both. The approach used was to determine the electrical excitability of the membrane by recording changes in membrane potential in response to imposed current.

The potential change recorded through one barrel of a double-barreled electrode while passing current through the other barrel is essentially ohmic. The recorded potential shifts abruptly to a new level during a current pulse and the potential change is proportional to the current intensity (fig. 7). The rise time of the potential change is very brief and suggests a circuit time constant of less than a millisecond. When the electrode pair was in the saline outside a muscle fiber the ratio of recorded potential change to applied current averaged 120 Kohm (range = 90-190 KO, n = 12). This represents a coupling resistance between the electrodes which depends on the tip dimensions of the electrodes and the conductivity of the surrounding medium. The average resistance recorded with the electrodes in a muscle fiber was 670 Kohm (range = 464-940 Kohm, penetrations of 6 fibers from each of 3 animals). It will be shown that a significant fraction of the measured resistance is due to interelectrode coupling which is greater when the electrodes are in cytoplasm than when they are in saline. The true input resistance of the muscle fiber must be less than that measured with double-barreled electrodes, but how much less cannot be determined without additional information. Electrical properties of muscle fibers measurements w i t h indepent current and voltage electrodes The voltage rise time recorded with a double-barreled electrode during current

CONTRACTION I N A FAST INSECT MUSCLE

289

A

Fig. 5 Mechanical responses of the DLM to nerve stimulation at 200 Hz, T = 3 5 ° C . In A the stimuli excited two fast axons. Increasing the stimulus intensity slightly (B) activated the slower axon. Further increasing the stimulus intensity (C) activated the last fast axon,

steps was suspiciously fast, suggesting as it did an extremely short membrane time constant. In order to minimize interelectrode coupling as a confounding parameter, the membrane response to imposed current was also examined using two independent electrodes, one for passing current and the other for recording potential. These experiments were done at 25°C. The distance between the tips of the electrodes was measured with an occular micrometer and ranged from 20 to 200 pm. Here i t will be assumed that this distance is short enough that the electrodes can be regarded as being at the same point in the muscle fiber. When the electrodes were separated by sim-

ilar distances but in adjacent fibers, no significant potential changes were recorded to imposed current; there is no electrical coupling between fibers. The input resistance to hyperpolarizing current or weak depolarizing current ranged from 50 to 350 Kohm (mean = 160 Kohm, 27 fibers from five animals). Data from one animal in which, for unknown reasons, the input resistance was discontinuously large (42&510 Kohm, 3 penetrations) have been excluded. Fiber damage during electrode insertion can be expected to reduce the input resistance. Therefore in analyzing the data, detailed attention has been given only to those fibers in the

290

ROBERT K. JOSEPHSON, DARRELL R. STOKES AND VICTOR CHEN

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higher end of the range (160-350 Kohm), it being felt that these are probably more representative of fibers in intact animals. The input resistance to hyperpolarizing current was constant over the entire current range tested (fig. 8). The membrane time constant, taken as the time to reach 84% of the asymptotic potential change (Hodgkin and Rushton, ’46), ranged from 0.8 to about 3 msec, 1.5 msec being a typical value. The voltage change with weak depolarizing currents is the mirror image of that to hyperpolarizing current of the same intensity (fig. 9). Outward current steps which depolarize the fiber more than %lo mV initiated a voltage transient or set of transients which usually reach a steady state in 10-20 msec. With strong depolarking currents the steady state input resistance drops (fig. 8); in some fibers the slope resistance becomes less than half its initial value as the fiber is progressively depolarized toward zero. The voltage transient at the onset of a depolarizing current pulse is a positive potential peak followed by a series of damped oscillations. The size of the initial peak and the size and frequency of following oscillations all increase with increasing outward current. Fig. 10 shows oscillations from one fiber in which these were unusually large. The interval between peaks of the oscillations is 6 9 msec. It might be noted that this is nearly the same as the interval between muscle contractions during singing ( 5 msec), especially when allowance is made for the lower muscle temperature during the imposed current measurements. In some fibers the potential rise during strong outward current is the mirror image of the potential change to inward current of equal intensity up to the peak of the initial voltage transient (fig. 11A). In such cases the initial peak could be explained simply as due to a delayed increase in membrane conductance (delayed rectification) which diminishes the potential change to imposed current. But in other fibers the potential at the initial peak is further from the resting potential than is the response to outward current of similar intensity after the same delay (fig. 11B). Here the potential displacement at the peak is greater than would be expected from the passive electrical properties of the membrane, indicating some, albeit weak, depolarizing electrogenesis.

29 1

CONTRACTION I N A FAST INSECT MUSCLE

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20 msec Fig. 7 Current-voltage relations of a muscle fiber as determined with a double barrel electrode. The closed circles were with the electrode in a muscle fiber. The open circles were collected immediately afterward with the electrode in the saline surrounding the fiber. The insets show the responses to depolarizing current (upper left) a n d to hyperpolarizing current (lower right); in each case the upper line indicating the zero potential level.

Hyperpolarizing a muscle fiber with injected current increases its electrical response to nerve stimulation and depolarizing the fiber decreases its nerve-initiated response (fig. 12). The muscle fibers are multiply-innervated and not space clamped, so the decline in electrical response with depolarization cannot simply be extrapolated to zero to obtain an equilibrium potential for the response (Burke and Ginsborg, '56). The relation between response amplitude and membrane potential is further complicated by the pronounced delayed rectification seen with these fibers. In analyzing an equivalent case Burke and Ginsborg concluded that the apparent reversal depolarization (the depolarization required to reduce the muscle response to zero, mea-

sured from the resting potential) could be as much as twice the true equilibrium depolarization. In figure 1 2 A both the points at high membrane potentials and those at very low membrane potentials extrapolate to zero response amplitude at about f 3 5 mV. For this fiber, whose resting potential was - 40 mV, the equilibrium potential of the muscle electrical response should lie somewhere between - 3 and 35 mV.

+

DISCUSSION

T h e nature of the muscle electrical response Stimulating the motornerve to the DLM initiates a short, often overshooting depolarization of the muscle fibers. Despite its spike-like appearance, available evidence

292

ROBERT K. JOSEPHSON, DARRELL R. STOKES AND VICTOR CHEN

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indicates that the muscle depolarization is primarily a synaptic potential. The muscle membrane shows little or no depolarizing electrogenesis in response to imposed current; with the exception of delayed rectification, the membrane often appears to be electrically inexcitable. In two other very fast, multiterminally-innervated muscles, the sound-producing muscle of the lobster

and that of the squirrelfish, injected current initiates only weak excitable responses while the depolarization to nerve stimulation appears to contain a major excitable component (Mendelson, '69; Gainer et al., '65). In both cases it was suggested that damage associated with electrode insertion may have diminished the membrane responsiveness to imposed current. While this

CONTRACTION I N A FAST INSECT MUSCLE

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1 I

293

is also a possibility for the katydid muscle, there is no evidence for it. The depolarization of katydid muscle following nerve stimulation does not have an inflection on its rising phase which might suggest an excitable response arising from a synaptic potential. Even strong hyperpolarizing current fails to dissociate the muscle response into a synaptic component and a triggered excitable membrane response (fig. 12; compare Cerf et al., '59). The muscle response to nerve stimulation is progressively reduced by depolarizing current and with strong depolarizing current the response may have about the same size and time course as the initial voltage transient initiated by the current pulse (fig. 12). The rising portion of the voltage transient appears to be determined largely by the passive electrical properties of the membrane, augmented in some instances (e.g., fig. 11B)by depolarizing electrogenesis,possibly due to an increase in sodium conductance (see Usherwood, '69, for discussion of graded electrogenesis in insect muscle). The subsequent decline of the voltage transient results from the onset of delayed rectification. The similarity between the voltage transient and the muscle response to nerve stimulation suggest that they are caused by a common mechanism, i.e., that the response to nerve input is a rapid depolarization due to synaptically-injected current, terminated in part by a delayed increase in the conductance of the muscle membrane. Electrical inexcitability in arthropod muscle is usually associated with slow contraction (e.g., Atwood, '73). It is therefore suprising to find it in an extraordinarily fast muscle. Operating with electricallyinexcitable membrane does offer one advantage; there is no refractory period. Even the graded excitable responses of insect muscle are followed by refractoriness. In locust leg muscle, partial refractoriness of the excitable response may last 25 msec (Cerf et al., '59). With an inexcitable membrane, refractoriness of the muscle membrane would not be a limit to response frequency. Katydids apparently use distributed synaptic potentials to activate their singing muscles. In this the muscles are like, for example, some leg muscles in locusts (Cerf et al., '59) or dragonfly larvae (Malpus, '68).

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KOBElRT K. JOSEPHSON, DARRELL R. STOKES AND VICTOR CHEN

L

L

-

I100nA

5 msec

Fig. 10 Potential oscillations during depolarizing current pulses. The upper trace indicates the zero potential level.

- -

I200 n A

10 msec

Fig. 1 1 Voltage transients at the onset of depolarizing current pulses. The response to hyperpolarizing current of the same intensity (dotted line) h a s been inverted and superimposed on that due to depolarizing current. A and B are from different fibers. The upper traces (only partially shown in A) are the zero potential levels.

Rapid activation and inactivation requires very brief synaptic potentials. Some features which reduce the time course of the synaptic potentials during singing are: (1) Operating at elevated temperature. A t room temperature the duration of the muscle

electrical response is about 4 msec; at 3 5 " C , the thoracic temperature during singing, the duration is reduced to about 2.5 msec. The increased temperature presumably accelerates transmitter release and subsequent inactivation. ( 2 ) A low resting mem-

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CONTRACTION IN A FAST INSECT MUSCLE

80

60

40

20 0

.

I

+20

0

-20

I

I

- 40

-60

I

-80

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-100

Membrane potential, mV

10 msec Fig. 12 The effect of membrane potential on the size of the response to nerve stimulation. The membrane potential was altered by imposed current pulses. The motorneurone-evoked depolarization of a fiber is plotted as a function of membrane potential in A . B, C, and D are sample responses from this fiber.

brane resistance and consequently a short (for muscle) membrane time constant (see below). The short time constant results in rapid decay of induced potential at the cessation of injected current. ( 3 ) A rapidly developing membrane conductance increase following depolarization (delayed rectification). This is probably a result of increasing potassium conductance (Cerf et al., '59). The conductance increase both shifts

the membrane potential toward a more negative level and further reduces the membrane time constant.

Passive electrical properties of the muscle membrane From cable theory (Hodgkin and Rushton, '46) the input resistance (Ro) of a long, cylindrical cell should be inversely related to the cell diameter (D). Specifically the input

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ROBERT K. JOSEPHSON, DARRELL R. STOKES A N D VICTOR C H E N

resistance should be proportional to the diameter raised to the negative three-halves power. The cells of the singing muscles are rather small, roughly 20-40 pm in diameter (Stokes et al., '75), yet their input resistance is surprisingly low, about 2 x 10.Y~.This is about the same as the input resistance of grasshopper extensor tibia muscle fibers (1.9 X lOzn) which have about twice the diameter of fibers from the singing muscle (Werman et al., '61, diameter from table 1 of Usherwood, '69). The low input resistance of the singing muscle fibers suggests that the membrane of these fibers has a low specific resistance. The input. resistance of a cylindrical cell depends on both the transverse membrane resistance and the internal longitudinal resistance: R o = 1/2 (rmriI1I2

Here rm is the resistance X unit length of the surface membrane and ri the longitudinal resistance per unit length of the cell interior. The usual method used to determine r, and ri is to measure the spatial decrement of the induced voltage during a long current pulse. From this the space constant (A) can be determined which is a function of rm and ri : A = (rm/ri)lIz

The size and mobility of katydid muscle cells made it sufficiently difficult to impale a cell with two electrodes, one for passing current and the other for measuring voltage, that we were discouraged from trying the multiple penetrations necessary to directly determine the space constants of these fibers. The internal longitudinal resistance of a cell can also be calculated from the specific resistivity (Ri) of the cytoplasm and the cross-sectional area of the cell (A): ri = R i / A

With some assumptions Ri can be calculated from the impedance measurements made with double-barreled electrodes. These assumptions are: (1) The true input resistance of the cells is that measured with two independent electrodes. (2) The cells sampled with two independent electrodes were similar to those sampled with doublebarreled electrodes so that the average cell with both methods had similar basic prop-

erties. ( 3 ) The DC coupling between barrels of a double-barreled electrode arises because the voltage electrode samples part of the three-dimensional voltage field surrounding the current electrode. The voltage gradient about the current electrode should be proportional to the specific resistivity of the cytoplasm (Eisenberg and Johnson, '70) so the coupling resistance should also be proportional to the cytoplasmic resistivity. (4) The coupling resistance with doublebarreled electrodes is in series with and additive to the membrane resistance. (5) With the exception of the resistance measured as electrode coupling, deviation from standard cable theory due to three-dimensional current spread about the current electrode (Eisenberg and Johnson, '70) is sufficiently small to be ignored in these approximations. The average input impedance recorded from 30 fibers with independent electrodes was 191 KR. This includes three suspiciously large values. The apparent input impedance recorded with double-barreled electrodes was 670 KO. The difference between these two values, 479 KO, presumably represents the coupling resistance between the two barrels when the doublebarreled electrodes are in cytoplasm. The coupling resistance with the double-barreled electrodes in saline was 120 KO. From the assumption that the coupling resistance is proportional to the specific resistivity of the medium surrounding the electrodes, the specific resistivity of the cytoplasm is four times greater than that of saline. The specific resistivity of saline was measured with a conductivity cell and found to be 62 Rcm. From this the specific resistivity of the cytoplasm (Ri) is 248 Ocm. Although this approach is quite indirect, the value obtained for Ri is similar to that obtained from other muscle fibers with more orthodox methods (e.g., crayfish 150 ncm, Law and Atwood, '71; frog semitendinous - 169 ncm, Hodgkin and Nakajima, '72; dragonfly leg muscles - 250 Ocm, Malpus, '68). Most of the resistance measurements with independent electrodes were made from muscle fibers i n the medial portion of the DLM, so the dimensions of medial band fibers will be used for further calculations. The average cross-sectional area of these fibers is 1.65 x lO-scm2 and the average

CONTRACTION I N A F A S T I N S E C T MUSCLE

circumference 1.67 x lO-zcm (Stokes et al., '75). From these values, the internal longitudinal resistance (ri) of a muscle fiber is 1.5 x 107 ncm-1, the transverse membrane resistance (r,) is 9.7 x 103 Rcm, and the fiber space constant (A) is 254 pm. This space constant is sufficiently short compared with the length of the fibers (approximately 4 mm) that it is appropriate to treat the fibers as cables of infinite length. The specific membrane resistance (R, .= r, x circumference) of these fibers is 162 ncm*. This is quite low; other reported values for insect muscle range from 530 Rcmz to 1.6 x 105 ncmz (table 1 in Usherwood, '69). It is presumably because of this low resistance that the membrane time constant ( T ~ )is short, about 1.5 msec. From this time constant the membrane capacity (Cm = T,/R,) is about 9 pF/crnz. Because of the low membrane resistance and associated short time constant, the voltage changes across the membrane to imposed currents are rapid. But there is a trade-off here; with a lowered membrane impedance the voltage change to a given current is reduced, hence more synaptic current must be generated for a given voltage change. This cost is not simply proportional to the specific membrane resistance as is the time constant. The synaptic potentials are short, essentially high frequency signals, and at high frequencies the membrane impedance depends more on the membrane capacitance than on its resistance. There is morphological evidence that the synapses are distributed along the length of the fibers (Stokes et al., '75). As a first approximation i t will be assumed that synaptic current is generated continuously along the fibers; the fiber can then be regarded as isopotential with no longitudinal current flow and voltage variation. In an isopotential fiber the synaptic current changes the potential across the surrounding membrane which has a transverse resistance R m and a parallel capacitance Cm. The impedance of the parallel RC circuit depends on the signal frequency, 250 Hz seems here a reasonable value (Katz, '66: p 99). With the membrane properties determined for singing muscle ( R m = 162 ncm2, Cm = 9 FF/cmz) the impedance for a 250 Hz signal is 65 Rcm2. Were the specific membrane resistance 5000 ncms, that

297

determined for a small frog sartorius muscle fiber (Hodgkin and Nakajima, '72), the impedance would be 71 flcm*. Thus changing the specific resistance by more than an order of magnitude changes the impedance to a 250 Hz signal by only 1 0 % .With either a high resistance or low resistance membrane the electrical impedance is low to high frequency signals. This means that for a given depolarization the synaptic current must be briefly high, and that because of the short effective space constant associated with the low impedance the synapses have to be rather close together to uniformly depolarize the membrane. To operate at high frequency is expensive in terms of synaptic current density and the number of synapses, but the cost is due more to the brevity of the responses than to the low DC resistance of the membrane. Facilitation of muscle activation The tension generated by an arthropod muscle fiber has been found to be monotonically related to the extent of membrane depolarization (e.g., Orkand, '62; Reuben et al., '67). On this basis one would expect facilitating contractile responses to be associated with facilitating electrical responses of the muscle membrane. In some singing muscle preparations there was clear mechanical facilitation to nerve stimulation while the associated membrane electrical responses defacilitated slightly and did not become noticeably longer. In figure 6A the stimulus frequency was such that the muscle relaxed almost completely between contractions. According to current dogma this indicates that the calcium concentration in the sarcoplasm falls below the mechanical threshold between each contraction, and hence that the facilitation cannot be explained as due to calcium accumulating in the sarcoplasm with a non-linear relation between calcium concentration and contractile activity. There appears to be facilitation of the excitation-contraction coupling process itself. One possibility is that there is facilitation of calcium release and resequestration by the sarcoplasmic reticulum during repetitive activity. Facilitating calcium influx during repetitive activity has been found in Aplysia neurones (Stinnakre and Tauc, '73), but this is associated with action potentials which become larger and longer. The performance of katydid muscle

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suggests that there may be facilitation of calcium movement without significant changes in electrical responses. LITERATURE C1TED Atwood, H. L. 1965 Characteristics of fibres in the extensor muscle of a crab. Comp. Biochem. Physiol., 14: 205-207. -- 1972 Crustacean muscle. I n : The Structure and Function of Muscle. Vol. I. G. H. Bourne, ed. Academic Press, New York a n d London. __ 1973 An attempt to account for the diversity of crustacean muscles. Amer. Zool., 13: 357378. Aulie, A , , and P. S . Enger 1969 The flight muscle in a bird with high wing-stroke frequency, the zebra finch. Physiol. Zool., 4 2 : 303-310. Bach-y-Rita, P . , and F. Ito 1966 I n vivo studies on fast and slow muscle fibers in c a t extraocular muscles. J . Gen. Physiol., 4 9 : 1177-1198. Becht, G., G. Hoyle and P. N. R. Usherwood 1960 Neuromuscular transmission in the coxal muscles of the cockroach. J . Insect Physiol., 4 : 191201. Benoit, P. 1962 Montage souple de microelectrodes intracellulaires, pour ]’etude des tissus contractiles. Comptes Rendus SOC. Biol., 156: 1465- 1466. Burke, W., and B. L. Ginsborg 1956 The action of neuromuscular transmitter on the slow fibre membrane. J . Physiol., 132: 5 9 9 4 1 0 . Cerf, J . A,, H. Grundfest, G. Hoyle a n d F. V. McCann 1959 The mechanism ofdual responsiveness i n muscle fibers of the grasshopper Romalen m i c r o p t e r u . J . Gen. Physiol., 4 3 : 377-395. Close, R. I., and Luff, A. R. 1974 Dynamic properties of inferior rectus muscles of the rat. J . Physiol., 2 3 6 : 2 5 S 2 7 0 . Ebashi, S . , a n d M. Endo 1968 Calcium ion a n d muscle contraction. Prog. Biophys. Mol. Biol., 18: 123-183. Eisenberg, R. S . , a n d E . A. Johnson 1970 Threedimensional electrical field problems in physiology. Prog. Biophys. Mol. Biol., 2 0 : 1 4 5 . Elder, H. Y . 1971 High fkequency muscles used i n sound production by a katydid. 11. Ultrastructure of the singing muscles. Biol. Bull., 141: 4 3 4 4 4 8 . Fatt, P., and B. Katz 1953 The electrical properties of crustacean muscle fibres. J . Physiol., 120: 171-204. Gainer, H., K . Kusano a n d R. F. Mathewson 1965 Electrophysiological and mechanical properties of squirrelfish sound-producing muscle. Comp. Biochem. Physiol., 1 4 : 6 6 1 4 7 1 . Hagiwara, S., S. Chichibu a n d N. Simpson 1968 Neuromuscular mechanisms of wing beat in hummingbirds. Z. vergl. Physiol., 60: 209-218. Heath, J . E., a n d R. K. Josephson 1970 Body temperature and singing i n the katydid, Neoconocephulus robustus (Orthoptera, Tettigoniidae). Biol. Bull., 138: 272-285. Henson, 0 .W., Jr. 1965 The activity a n d function of the middle-ear muscles i n echo-locating bats. J . Physiol., 180: 871-887. Hodgkin, A. L., a n d S. Nakajima 1972 The effect ofdiameter on the electrical constants offrog skeletal muscle fibres. J . Physiol., 221 : 105-120.

Hodgkin, A. L., a n d W. A. H. Rushton 1946 The electrical constants of a crustacean n e m e fibre. Proc. Roy. SOC.Lond. B., 133: 4 4 4 4 7 9 . Hoyle, G. 1954 Changes in the blood potassium concentration of the African migratory locust ( Lo c u s tu migrntoriu migrntorioides R. & F.1 during food deprivation, and the effect on neuromuscular activity. J . Exp. Biol., 31 : 260-270. Hoyle, G., and M. Burrows 1973 Correlated physiological and ultrastructural studies on specialized muscles. IIIa. Neuromuscular physiology of the power-stroke muscle of the swimming leg of Portunccs s u n g u i n o l e n t u s . J . Exp. Zool., 185. 8396. Huxley, A. F. 1971 The activation of striated muscle a n d its mechanical response. Proc. Roy. SOC. Lond. B., 178: 1-27. Josephson, R. K. 1973 Contraction kinetics of the fast muscles used in singing by a katydid. J . Exp. Biol., 5 9 : 781-801. Josephson, R. K., and R. C. Halverson 1971 High frequency muscles used in sound production by a katydid. I . Organization of the motor system. Biol. Bull., 141: 411-433. Katz, B. 1966 Nerue, M u s c l e u n d S y n a p s e . McGraw Hill, New York. Law, P. K., a n d H. L. Atwood 1971 Sarcoplasmic resistivity of crayfish muscle fibers. Canadian J . Zool., 4 9 : 1063-1064. Malpus, C. M. 1968 Electrical responses of muscle fibres of dragonfly larvae in relation to those of other insects a n d ofcrustaceans. J. Insect Physiol., 1 4 : 128.51301. Mendelson, M. 1967 A simple method of fabricating double-barrelled micropipette electrodes. J . Sci. Instrum., 44: 5 4 S 5 5 0 . 1969 Electrical and mechanical characteristics of a very fast lobster muscle. J. Cell Biol., 4 2 : 548-563. Neville, A. C . , and T. Weis-Fogh 1963 The effect of temperature on locust flight muscle. J . Exp. Biol., 40: 111-121. Orkand, R. K. 1962 The relation between membrane potential and contraction i n single crayfish muscle fibres. J . Physiol., 161 : 143-159. Ozeki, M. 1969 Crayfish muscle fiber: spike electrogenesis in fibers with long sarcomeres. Science, 163: 8%83. P a r n a s , I., and H. L. Atwood 1966 Phasic a n d tonic neuromuscular systems in the abdominal extensor muscles of the crayfish and rock lobster. Comp. Biochem. Physiol., 1 8 : 701-723. Reuben, J . P., P. W. Brandt, H. Garcia and H . Grundfest 1967 Excitation-contraction coupling i n crayfish. Amer. Zool., 7: 62%645. Stinnakre, J., and L. Tauc 1973 Calcium influx in active A p l y s i a neurones detected by injected aequorin. Nature New Biology, 242: 113-115. Stokes, D. R., R. K. Josephson and R. B. Price 1975 Structural a n d functional heterogeneity i n a n insect muscle. J. Exp. Zool., in press. Usherwood, P. N. R. 1962 The nature of “slow” and “fast” contractions in the coxal muscles of the cockroach. J . Insect Physiol., 8 : 31-52. 1968 A critical study of the evidence for peripheral inhibitory axons in insects. J . Exp. Biol., 4 9 : 201-222. 1969 Electrochemistry of insect muscle. Adv. Insect Physiol., 6: 205-278.

CONTRACTION IN A FAST INSECT MUSCLE Weber, A , , and J. Murray 1973 Molecular control mechanisms in muscle contraction. Physiol. Rev., 5 3 : 612-673. Werman, R., F. V. McCann and H. Grundfest 1961

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Graded and all-or-none electrogenesis in arthropod muscle. I. The effects of alkali-earth cations on the neuromuscular system of Romctletr microptcrcz. J. Gen. Physiol.,44: 97S995.