Cell Culture in Neurobiology

Neurophysiology > Cellular Biology of Neurons > Cellular Interactions

Email a friend
Contact the editor about this article
How to cite

Gerald D. Fischbach, Phillip G. Nelson

10.1002/cphy.cp010120

Source: Supplement 1: Handbook of Physiology, The Nervous System, Cellular Biology of Neurons

Originally published: 1977

Published online: January 2011

Full Article on Wiley Online Library

Abstract
Images
References

Abstract

The sections in this article are:

1 Introduction

1.1 Primary Cell Cultures

1.2 Cell Lines

1.3 Muscles as Models

2 Techniques

2.1 Primary Cultures

2.2 Cell Lines

3 Cell Form

3.1 Distinctive Cell Shapes

3.2 Development of Cell Form

3.3 Growth Cones

3.4 Factors Affecting Process Formation

4 Resting Membrane Potential

5 Action Potential

5.1 Ionic Determinants

5.2 Developmental Aspects

6 Synaptic Transmission

6.1 Nerve‐Muscle Synapses

6.2 Synapses Between Neurons

6.3 Chemosensitivity of Neurons

6.4 Transmitter Metabolism

6.5 Factors Affecting Synapse Formation

7 Summary

	Figure 1. Rat spinal cord and attached spinal ganglion explant culture after 46 days in vitro. Arrows point to spinal ganglion cells. Tissue organization, including dorsal and ventral roots, is preserved to a remarkable degree. Note that fibers originating in the anterior horn or attached ganglion tend to follow a fairly straight course. Only myelinated fibers are seen in this bright‐field view From Sobkowicz et al. 323
	Figure 2. Dissociation of a 7‐day embryonic chick spinal cord; at this age sensory ganglia are located within the cartilaginous spinal canal. A: sensory ganglia remain attached to the spinal cord when it is removed. B: same segment shown in A after “stripping’ the dorsal root ganglion by removing the meninges. C: minced fragments of the spinal cord. D: after incubating fragments in 0.1% trypsin in Pucks (divalent cation‐free) saline, they are resuspended in complete medium, triturated, and filtered. E: isolated, spherical cells. Note variation in cell diameter: vertical bar in B represents 1 mm and applies to A‐C; horizontal bar in E, 50 μm.
	Figure 3. Elimination of “background’ cells. Skeletal muscle fibers in a control (A) 12‐day culture and in a sister culture (α) treated with cytosine arabinoside (araC; 10⁻⁵ M) for 48 h beginning on the 3rd day. Inset in α is a typical hypolemmal muscle nucleus. Spinal cord neurons in a control (B) and in a sister culture (b) treated with araC. Sympathetic ganglion cells grown in media containing HCO₃⁻ and exposed to CO₂ (C) and in media without HCO₃⁻ and exposed to air (c). Horizontal bars, 50 μm in each panel. Note the absence of background, fibroblast‐like cells in a‐c A and α from Fischbach & Cohen 93; B and b from Fischbach 91; C and c from P. Patterson, unpublished observations
	Figure 4. Separation of dissociated 7‐day chick spinal cord cells by velocity sedimentation (see text). Interference contrast micrographs illustrate the wide range in cell size in the initial suspension (upper), small cells in trailing fractions (middle), and large cells in leading fractions (lower). From D. K. Berg and G. D. Fischbach, unpublished observations
	Figure 5. Neuroblastoma cells clone N‐18. A: log phase. B: stationary phase — grown in the absence of serum for 4 days. C: aminopterin‐selected (10⁻⁶ M) cells grown in the presence of serum From Peacock et al. 251
	Figure 6. Silver‐impregnated spinal cord cells. A and B illustrate the range in size and shape of neurons. Compare with the characteristically bipolar sensory ganglion cell (C) grown under identical conditions. Horizontal bar, 50 μm From Fischbach & Dichter 95
	Figure 7. Isolated sympathetic ganglion cell: A, at time 0 (18 h after plating); B, at 60 min; C, at 120 min; D, at 240 min; E, at 360 min; F, at 500 min. A‐C same scale, horizontal bar 78 μm, D‐F same scale, horizontal bars 80 μm. Note that at the end of the sequence all growing tips are approximately equidistant from the cell body and also that branch points are apparently stable.
	Figure 8. Polarity of processes. A: sensory ganglion cell with one process issuing from the cell body after 4 days in culture. From Nakai 222.] Note the “recurrent collateral’ near the cell body. B: spinal cord neuron in a 3‐wk culture with one prominent, long, relatively unbranched process. Cell body inked in. [From Fischbach 90. Copyright 1970 by the American Association for the Advancement of Science
	Figure 9. Growth of chick embryonic neurites in culture plotted from time‐lapse film records. A: control; small dots refer to a sensory ganglion cell and larger dots to a neuron isolated from the midbrain. B: neurites severed from their cell body at time 0 Adapted from Hughes 143
	Figure 10. Scanning electron micrographs of chick sensory ganglion cell growth cones 1 day after plating. Note the fine microspikes in A and the elaborate “undulating’ membrane in B. A, × 2,000; B, × 17,000. Micrographs provided by J. Jabaily
	Figure 11. Ultrastructure of a chick sensory ganglion cell growth cone. A filamentous network (FN) which extends into the microspikes is evident beneath the surface membrane. Note that microtubules (MT) appear to end at the base of the growth cone and that neurofilaments (NF) are centrally located. Tubulovesicular profiles of the smooth endoplasmic reticulum (SER) are abundant and appear at the base of one microspike. V, D, and C are smooth, dense‐core, and coated vesicles, respectively; MC is a mitochondrion From Yamada et al. 368
	Figure 12. Selective growth of sympathetic ganglion (sg) neurites toward nearby fragments of vas deferens (vd) rather than kidney (k); 5 days in vitro. Horizontal bar, 400 μm From Chamley et al. 46
	Figure 13. Membrane potential (A) and the relation between membrane potential and [K⁺]₀(B) of rat myotubes as a function of age. In A, • refers to multinucleated myotubes and Δ to mononucleated myoblasts. Vertical bars, ± SE. The number of cells at each point is indicated. In B, 4 myotubes of the indicated ages were examined. In each case the cultures were perfused with a continuous K⁺ gradient between 5.3 mM and 148 mM which was completed in 3 min. The relation between membrane potential and log [K⁺]₀ predicted by the Nernst equation is indicated by the line labeled E_K. Continuous lines through each set of points are predicted from the constant‐field equation (Eq. 1 in text) assuming [Na]_i = 13 mM and the P_Na⁺‐to‐P_K⁺ ratios are as indicated at each day From Ritchie & Fambrough 283
	Figure 14. Action potentials evoked in spinal cord (A and C) and sensory ganglion (B and D) cells. Duration of current pulses indicated by bars above each trace; dorsal root ganglion (DRG) spikes are prolonged and are marked by a plateau on the falling phase. Many spinal cord neurons fire repetitively during a sustained depolarization (C), whereas most DRG cells rapidly adapt (D). The resting potential of cells in A‐C was 50–55 mV.
	Figure 15. Action potentials conducted along a process. All records obtained from a microelectrode located in the cell body shown in A. Spikes were evoked by extracellular stimulation (frames 2–4) as shown and by intracellular stimulation (frame 1). Stimulus artifacts appear as breaks in the records. Note the inflection on the rise of conducted spikes. In B‐4, superimposed traces show that hyperpolarization (inward current pulse not shown) accentuates the inflection and ultimately blocks the second component. Calibration pulse, 20 mV, 1 ms. Vertical bar in A, 50 μm From Fischbach & Dichter 95
	Figure 16. Presumptive glial cells. A and B: two cells found to be electrically inexcitable. Current‐voltage relation of the cell shown in B is plotted in C. Sample traces are shown in inset. Note that the relation remains linear over a wide range of membrane potential.
	Figure 17. Ca²⁺ spikes in sensory neurons. A: relation between overshoot of the action potential evoked at the cell body and external Ca²⁺. Insets show representative traces recorded in the presence of TTX (10⁻⁷ g/ml) and 1.8 mM Ca²⁺ (trace 1) and 18 mM Ca²⁺ (trace 2). B: effect of Na⁺ on conducted spikes. Two cells (α and b) were bathed in Na⁺‐free media. In each case stimulation (arrows) of a process 300–400 μm away from the cell body failed to evoke a conducted spike (trace 1). An action potential was evoked via an intrasoma electrode in cell b. Immediately before trace 2, a 50 μm tip, NaCl‐filled electrode was lowered adjacent to the process. Identical extracellular stimuli now evoked conducted spikes in α and b. The axon spike in α invaded the cell body, whereas the one in b did not. Trace 3 in b was recorded after the NaCl electrode was removed. Note in b the decrease in duration of the soma spike in the Na⁺ atmosphere From Dichter & Fischbach 370
	Figure 18. Action potential mechanisms in chick (A) and L‐6 (B) myotubes. A: in chick fibers, the sharp spike observed in normal saline (1) is abolished in Na⁺‐free (substituted with sucrose) media (2); a small approximately 50‐ms active response remains (wedge) which is abolished by Co²⁺ (10 mM) (3). The prolonged active response (arrows) which is not affected by removing Na⁺ or by addition of Co²⁺ is presumably due to an outward Cl⁻ current. The stimulus duration is indicated by a bar below each trace. From Fuduka 102.] B: in normal saline the sharp spike of L‐6 myotubes (evoked by anode‐break excitation) is followed by a prolonged active response (1). The sharp spike is absent, but the late response is increased in size and duration in Na⁺‐free saline containing 10 mM Ca²⁺ (2). (Normal saline contains 1.8 mM Ca²⁺.) [From Kidokoro 164
	Figure 19. Changes in action potential generation in two cell lines as they “differentiate’ in vitro. A: L‐6 muscle cells. A small active response evoked in L‐6 mononucleated cell (myoblast) following a hyperpolarizing current pulse is contrasted with a large prolonged spike in a multinucleated (fused) myotube. In each pair, current is displayed in the upper trace and membrane potential in the lower trace. From Kidokoro 164.] B: neuroblastoma cells (clone N‐18) in log phase (upper record), stationary phase (middle record), and selected in the presence of aminopterin (lower record). [From Peacock et al. 251.] The neuroblastoma spikes were recorded from the cells in Fig. 5 which are indicated by the blurred image of a microelectrode
	Figure 20. Action potentials in neuroblastoma × L cell hybrids. Cells from two clones derived from the same hybridization are shown in A and B. A appears more neuronal and B more fibroblastic. The response of each cell type to depolarizing currents is shown in adjacent records. Cells in the neuronlike clone generate spikes, whereas those in the fibroblastlike clone do not. Both clones were grown in the presence of aminopterin for 10 days. Calibration pulse, 10 mV From Peacock et al. 250
	Figure 21. A multipolar chick spinal cord neuron that has settled between muscle fibers on the collagen substratum. The neuron was dissociated from a 7‐day cord and plated about 48 h earlier. Note the microspike that extends from an expanded growth cone (gc) to contact a muscle fiber.
	Figure 22. Neuromuscular synaptic physiology. A: functional contact — a depolarizing synaptic potential (upper trace) follows an action potential in a nearby neuron evoked by intracellular stimulation (current pulse not shown). Bars, 3.5 and 50 mV for muscle and neuron records, respectively, and 10 ms for both. B: superimposed traces show the change in end‐plate potential (EPP) amplitude as the muscle membrane potential (V_m) was altered to the values shown at the left of each record by steady currents injected through a second intracellular microelectrode; calibration pulse, 5 mV, 5 ms. Each point in the graph represents the mean amplitude of 5–10 EPP's. C: ACh response (arrow) evoked by a 1‐ms pulse of ACh precisely mimics a spontaneous synaptic potential recorded on the same trace; calibration, 5 mV, 10 ms. D: reversible reduction in EPP size by d‐tubocurare (10⁻⁷ g/ml); record at extreme right obtained 30 min after removal of the drug; bars, 15 mV, 15 ms. E: miniature end‐plate potentials (MEPP's) recorded on contiguous segments of moving film; bars, 5 mV, 2 s. F: variation in amplitude (and one failure) of successive EPP's in a series evoked at a rate of 1/s, despite a constant stimulus (action potential) in the innervating neuron (upper traces); calibration pulse, 5 mV, 5 ms A, B, D, and F from Fischbach 91; C from Fischbach & Cohen 93
	Figure 23. A site of transmitter release identified by focal depolarization in the presence of TTX (see text). Interference contrast illumination of a chick muscle cell contacted by a neurite from a nearby spinal cord explant. Synaptic potentials were evoked by extracellular stimulation at α but not at b or elsewhere along the process which extends along the edge of the fiber From Fischbach et al. 372
	Figure 24. Appearance of ACh receptors in muscle cells. A: increase in α‐bungarotoxin (α‐BGT) binding in control (▪) and in fusion‐arrested (•) chick muscle cultures (see text). ▴, fusion index (percent of nuclei incorporated into myotubes). Δ, fusion index in low‐Ca²⁺ media. Note that the increase in toxin binding lags behind the fusion index in control cultures and is even further delayed when fusion is blocked. From Patterson & Prives 244.] B: elongated mononucleated muscles from rat muscle grown in standard media for 48 h. The oscilloscope record shows an ACh response (current on lower trace) recorded in cell 1. Horizontal bar, 100 μm. [From Fambrough & Rash 89.] C: autoradiographs of freshly dissociated (without the use of enzymes) mononucleated cells from chick muscle exposed in suspension to [¹²⁵I]α‐BGT. Interference contrast and bright‐field views. Horizontal bar, 10 μm. [From Smilowitz et al. 322
	Figure 25. Synthesis and degradation of ACh receptors by cultured muscle fibers. A: following a saturating exposure to unlabeled α‐bungarotoxin (α‐BGT). The appearance of new receptors was determined by [¹²⁵I]α‐BGT binding (•). ○, cultures treated as above but pretreated for 3 h with cycloheximide; Δ, cultures preincubated in dinitrophenol plus iodoacetate. From Harzell & Fambrough 128.] B: release of radioactivity into the medium from chick or rat muscle fibers saturated with [¹²⁵I]α‐BGT. Release, expressed as a fraction of that initially bound, is a first‐order process with a half‐time of about 20 h. [From Devreotes & Fambrough 73
	Figure 26. Histograms of ACh sensitivity of uninnervated (A) and innervated (B) muscle fibers in spinal cord muscle cell cultures. From Fischbach and Cohen 93.] In (C) active and inactive uninnervated fibers are compared. No neurons were present. Active fibers stimulated intermittently for 2–3 days at 10/sec (\ \ \ \) were less sensitive than inactive fibers grown in the presence of TTX (/ / / /). [From Cohen & Fischbach 53. Copyright 1973 by the American Association for the Advancement of Science
	Figure 27. Relative peaks of ACh sensitivity on uninnervated (A) and innervated (B) muscle fiber; ordinate, mV/nC. In A, bars indicate position of hypolemmal muscle nuclei. [From Fischbach & Cohen 93.] B: a hot spot located exactly at a site of transmitter release. Inset, synaptic potentials evoked by focal depolarization at the termination of a nerve process. ACh was iontophoresed at the points indicated in the tracing (prepared from a photograph), and the sensitivities at each point are shown in register below. Bars, 5 mV, 5 ms. From S. A. Cohen and G. D. Fischbach, unpublished observations
	Figure 28. Tubulovesicular network of cultured muscle fibers. In A note the openings to the exterior and also that some tubules form junctions with cisternae of the sarcoplasmic reticulum (arrow). Horizontal bar, 0.25 μm. B: unstained section of a muscle fiber exposed to horseradish peroxidase prior to fixation. The reaction product outlines areas continuous with the extracellular space. N, nucleus; G, Golgi; T, tubules; S, sarcoplasmic reticulum. Micrographs provided by M. Henkart
	Figure 29. Spontaneously occurring synaptic potentials in dissociated chick spinal cord cells. A and B: superimposed traces show depolarizing potentials (EPSP's) in one cell (A), hyperpolarizing potentials (IPSP's) in another (B, upper record), and both in a third cell (B, lower record). Vertical bar, 10 mV for A, 12 mV for B. Horizontal bar, 5 ms for A, 10 ms for B, upper trace, 20 ms for B, lower trace. C: a continuous record shows regularly occurring large EPSP's that often trigger action potentials. Note the few small PSP's. D: Small PSP's recorded in the presence of TTX (10⁻⁷ g/ml). Vertical bar, 10 mV for C, 5 mV for D. Horizontal bar, 1 s for C, 5 s for D From Fischbach & Dichter 95
	Figure 30. Stimulus‐evoked synaptic potentials in mouse spinal cord‐sensory ganglion cell cultures. A: monophasic EPSP (trace 3) evoked in one spinal cord neuron by stimulation (current in trace 1) of another spinal cord neuron (trace 2). B: action potential in a sensory neuron (evoked by a brief current pulse — not shown); trace 1 is followed by a small variable EPSP in a spinal cord cell (superimposed traces in 2). A larger less variable EPSP evoked by another sensory neuron in another spinal cord cell is shown in trace 3. C: complex responses in spinal cord neurons. An outward current plus (trace 3) evokes three spikes and is followed by a long‐lasting barrage of spikes and PSP's in the same (trace 2) and in an adjacent (trace 1) neuron.
	Figure 31. Electron micrographs of synaptic structures in chick spinal cord (A) and rat brain (B and C) cell cultures. In each, note the accumulations of small clear vesicles, the larger dense‐core vesicles, and increased density or thickening of the synaptic membranes. A, Several synaptic profiles appear along a single process. × 18,000. C: a synapse contacts a small process which may be a “spine.’ B and C, × 32,000. A, from Fischbach & Dichter 95; B and C provided by E. Neale
	Figure 32. Stimulation of a single presynaptic ending on a chick spinal cord neuron. Successive brief negative current pulses, delivered through a 2‐μm‐tipped NaCl‐filled pipette placed exactly over the bouton indicated by the arrow, evoked the synaptic potentials shown at right (traced from photographs). The response disappeared when the stimulating electrode was moved about 5 μm in any direction. Note that the incoming nerve processes pass out of the plane of focus. The postsynaptic nerve cell body overlies a muscle fiber. Calibration bars, 5 mV, 5 ms.
	Figure 33. Drug responses. A: ACh potentials in a sympathetic ganglion cell. The 20‐nA iontophoretic pulse is shown in the second trace; suprathreshold and subthreshold responses are shown in a and b. B: glutamate potential in a spinal cord neuron; iontophoretic pulse, 30 nA. This response is slower than EPSP's recorded in the same cell (trace 3). C: GABA potentials in a sympathetic ganglion cell. D: GABA potentials in a spinal cord neuron. A 25‐nA pulse (duration indicated by bar) was repeated as the membrane potential was shifted. The response reverses in polarity between −70 and −50 mV A and C from Obata 233; B and D from B. R. Ransom and P. G. Nelson, unpublished observations
	Figure 34. Nonuniform distribution of chemoreceptors on a mouse spinal cord neuron (A) and a neuroblastoma cell (B) about 4 wk after plating. A: penwriter records of glutamate responses at the sites indicated. Note the relatively large and rapid response at lower left. The same neuron impregnated with silver is shown in the inset. From B. R. Ransom and P. G. Nelson, unpublished observations.) B: ACh produces a qualitatively different response at different sites — a depolarization over the soma, a hyperpolarization at relatively distant sites along processes, and a biphasic response at intermediate points. (From Peacock et al. 1973
	Figure 35. A: increase in synthesis of catecholamines (norepinephrine and dopamine) from [³H]tyrosine by rat sympathetic ganglion neurons with time in culture. Note the initial log of 7–8 days followed by a 50‐fold increase. This pattern is markedly different from the increase in total accumulation of radioactivity measured in the same cultures (B) From Mains & Patterson 196
	Figure 36. Cholinergic synapses between sympathetic ganglion cells. A: simultaneous recordings from two sympathetic neurons in which each spike in one (the “driver,’ middle trace) is followed by a depolarizing synaptic potential in the other (the “follower,’ upper trace). Some PSP's in the follower are large enough to trigger spikes. The first spike in the driver cell is evoked by a brief pulse of outward current (lower trace). The other spikes arise spontaneously. B: the same cells in the presence of 50 μM hexamethonium, a nicotinic cholinergic antagonist. All spontaneous PSP's are abolished, and the evoked spike in the driver neuron is no longer followed by a PSP in the follower. C: the same cells after washing out the hexamethonium. Calibration bars: 40 mV, 40 ms. From O. MacLeish, P. H. O'Lague, E. J. Furshpan, and D. D. Potter, unpublished observations

Figure 1.

Rat spinal cord and attached spinal ganglion explant culture after 46 days in vitro. Arrows point to spinal ganglion cells. Tissue organization, including dorsal and ventral roots, is preserved to a remarkable degree. Note that fibers originating in the anterior horn or attached ganglion tend to follow a fairly straight course. Only myelinated fibers are seen in this bright‐field view

From Sobkowicz et al. 323

Figure 2.

Dissociation of a 7‐day embryonic chick spinal cord; at this age sensory ganglia are located within the cartilaginous spinal canal. A: sensory ganglia remain attached to the spinal cord when it is removed. B: same segment shown in A after “stripping’ the dorsal root ganglion by removing the meninges. C: minced fragments of the spinal cord. D: after incubating fragments in 0.1% trypsin in Pucks (divalent cation‐free) saline, they are resuspended in complete medium, triturated, and filtered. E: isolated, spherical cells. Note variation in cell diameter: vertical bar in B represents 1 mm and applies to A‐C; horizontal bar in E, 50 μm.

Figure 3.

Elimination of “background’ cells. Skeletal muscle fibers in a control (A) 12‐day culture and in a sister culture (α) treated with cytosine arabinoside (araC; 10⁻⁵ M) for 48 h beginning on the 3rd day. Inset in α is a typical hypolemmal muscle nucleus. Spinal cord neurons in a control (B) and in a sister culture (b) treated with araC. Sympathetic ganglion cells grown in media containing HCO₃⁻ and exposed to CO₂ (C) and in media without HCO₃⁻ and exposed to air (c). Horizontal bars, 50 μm in each panel. Note the absence of background, fibroblast‐like cells in a‐c

A and α from Fischbach & Cohen 93; B and b from Fischbach 91; C and c from P. Patterson, unpublished observations

Figure 4.

Separation of dissociated 7‐day chick spinal cord cells by velocity sedimentation (see text). Interference contrast micrographs illustrate the wide range in cell size in the initial suspension (upper), small cells in trailing fractions (middle), and large cells in leading fractions (lower).

From D. K. Berg and G. D. Fischbach, unpublished observations

Figure 5.

Neuroblastoma cells clone N‐18. A: log phase. B: stationary phase — grown in the absence of serum for 4 days. C: aminopterin‐selected (10⁻⁶ M) cells grown in the presence of serum

From Peacock et al. 251

Figure 6.

Silver‐impregnated spinal cord cells. A and B illustrate the range in size and shape of neurons. Compare with the characteristically bipolar sensory ganglion cell (C) grown under identical conditions. Horizontal bar, 50 μm

From Fischbach & Dichter 95

Figure 7.

Isolated sympathetic ganglion cell: A, at time 0 (18 h after plating); B, at 60 min; C, at 120 min; D, at 240 min; E, at 360 min; F, at 500 min. A‐C same scale, horizontal bar 78 μm, D‐F same scale, horizontal bars 80 μm. Note that at the end of the sequence all growing tips are approximately equidistant from the cell body and also that branch points are apparently stable.

Figure 8.

Polarity of processes. A: sensory ganglion cell with one process issuing from the cell body after 4 days in culture.

From Nakai 222.] Note the “recurrent collateral’ near the cell body. B: spinal cord neuron in a 3‐wk culture with one prominent, long, relatively unbranched process. Cell body inked in. [From Fischbach 90. Copyright 1970 by the American Association for the Advancement of Science

Figure 9.

Growth of chick embryonic neurites in culture plotted from time‐lapse film records. A: control; small dots refer to a sensory ganglion cell and larger dots to a neuron isolated from the midbrain. B: neurites severed from their cell body at time 0

Adapted from Hughes 143

Figure 10.

Scanning electron micrographs of chick sensory ganglion cell growth cones 1 day after plating. Note the fine microspikes in A and the elaborate “undulating’ membrane in B. A, × 2,000; B, × 17,000.

Micrographs provided by J. Jabaily

Figure 11.

Ultrastructure of a chick sensory ganglion cell growth cone. A filamentous network (FN) which extends into the microspikes is evident beneath the surface membrane. Note that microtubules (MT) appear to end at the base of the growth cone and that neurofilaments (NF) are centrally located. Tubulovesicular profiles of the smooth endoplasmic reticulum (SER) are abundant and appear at the base of one microspike. V, D, and C are smooth, dense‐core, and coated vesicles, respectively; MC is a mitochondrion

From Yamada et al. 368

Figure 12.

Selective growth of sympathetic ganglion (sg) neurites toward nearby fragments of vas deferens (vd) rather than kidney (k); 5 days in vitro. Horizontal bar, 400 μm

From Chamley et al. 46

Figure 13.

Membrane potential (A) and the relation between membrane potential and [K⁺]₀(B) of rat myotubes as a function of age. In A, • refers to multinucleated myotubes and Δ to mononucleated myoblasts. Vertical bars, ± SE. The number of cells at each point is indicated. In B, 4 myotubes of the indicated ages were examined. In each case the cultures were perfused with a continuous K⁺ gradient between 5.3 mM and 148 mM which was completed in 3 min. The relation between membrane potential and log [K⁺]₀ predicted by the Nernst equation is indicated by the line labeled E_K. Continuous lines through each set of points are predicted from the constant‐field equation (Eq. 1 in text) assuming [Na]_i = 13 mM and the P_Na⁺‐to‐P_K⁺ ratios are as indicated at each day

From Ritchie & Fambrough 283

Figure 14.

Action potentials evoked in spinal cord (A and C) and sensory ganglion (B and D) cells. Duration of current pulses indicated by bars above each trace; dorsal root ganglion (DRG) spikes are prolonged and are marked by a plateau on the falling phase. Many spinal cord neurons fire repetitively during a sustained depolarization (C), whereas most DRG cells rapidly adapt (D). The resting potential of cells in A‐C was 50–55 mV.

Figure 15.

Action potentials conducted along a process. All records obtained from a microelectrode located in the cell body shown in A. Spikes were evoked by extracellular stimulation (frames 2–4) as shown and by intracellular stimulation (frame 1). Stimulus artifacts appear as breaks in the records. Note the inflection on the rise of conducted spikes. In B‐4, superimposed traces show that hyperpolarization (inward current pulse not shown) accentuates the inflection and ultimately blocks the second component. Calibration pulse, 20 mV, 1 ms. Vertical bar in A, 50 μm

From Fischbach & Dichter 95

Figure 16.

Presumptive glial cells. A and B: two cells found to be electrically inexcitable. Current‐voltage relation of the cell shown in B is plotted in C. Sample traces are shown in inset. Note that the relation remains linear over a wide range of membrane potential.

Figure 17.

Ca²⁺ spikes in sensory neurons. A: relation between overshoot of the action potential evoked at the cell body and external Ca²⁺. Insets show representative traces recorded in the presence of TTX (10⁻⁷ g/ml) and 1.8 mM Ca²⁺ (trace 1) and 18 mM Ca²⁺ (trace 2). B: effect of Na⁺ on conducted spikes. Two cells (α and b) were bathed in Na⁺‐free media. In each case stimulation (arrows) of a process 300–400 μm away from the cell body failed to evoke a conducted spike (trace 1). An action potential was evoked via an intrasoma electrode in cell b. Immediately before trace 2, a 50 μm tip, NaCl‐filled electrode was lowered adjacent to the process. Identical extracellular stimuli now evoked conducted spikes in α and b. The axon spike in α invaded the cell body, whereas the one in b did not. Trace 3 in b was recorded after the NaCl electrode was removed. Note in b the decrease in duration of the soma spike in the Na⁺ atmosphere

From Dichter & Fischbach 370

Figure 18.

Action potential mechanisms in chick (A) and L‐6 (B) myotubes. A: in chick fibers, the sharp spike observed in normal saline (1) is abolished in Na⁺‐free (substituted with sucrose) media (2); a small approximately 50‐ms active response remains (wedge) which is abolished by Co²⁺ (10 mM) (3). The prolonged active response (arrows) which is not affected by removing Na⁺ or by addition of Co²⁺ is presumably due to an outward Cl⁻ current. The stimulus duration is indicated by a bar below each trace.

From Fuduka 102.] B: in normal saline the sharp spike of L‐6 myotubes (evoked by anode‐break excitation) is followed by a prolonged active response (1). The sharp spike is absent, but the late response is increased in size and duration in Na⁺‐free saline containing 10 mM Ca²⁺ (2). (Normal saline contains 1.8 mM Ca²⁺.) [From Kidokoro 164

Figure 19.

Changes in action potential generation in two cell lines as they “differentiate’ in vitro. A: L‐6 muscle cells. A small active response evoked in L‐6 mononucleated cell (myoblast) following a hyperpolarizing current pulse is contrasted with a large prolonged spike in a multinucleated (fused) myotube. In each pair, current is displayed in the upper trace and membrane potential in the lower trace.

From Kidokoro 164.] B: neuroblastoma cells (clone N‐18) in log phase (upper record), stationary phase (middle record), and selected in the presence of aminopterin (lower record). [From Peacock et al. 251.] The neuroblastoma spikes were recorded from the cells in Fig. 5 which are indicated by the blurred image of a microelectrode

Figure 20.

Action potentials in neuroblastoma × L cell hybrids. Cells from two clones derived from the same hybridization are shown in A and B. A appears more neuronal and B more fibroblastic. The response of each cell type to depolarizing currents is shown in adjacent records. Cells in the neuronlike clone generate spikes, whereas those in the fibroblastlike clone do not. Both clones were grown in the presence of aminopterin for 10 days. Calibration pulse, 10 mV

From Peacock et al. 250

Figure 21.

A multipolar chick spinal cord neuron that has settled between muscle fibers on the collagen substratum. The neuron was dissociated from a 7‐day cord and plated about 48 h earlier. Note the microspike that extends from an expanded growth cone (gc) to contact a muscle fiber.

Figure 22.

Neuromuscular synaptic physiology. A: functional contact — a depolarizing synaptic potential (upper trace) follows an action potential in a nearby neuron evoked by intracellular stimulation (current pulse not shown). Bars, 3.5 and 50 mV for muscle and neuron records, respectively, and 10 ms for both. B: superimposed traces show the change in end‐plate potential (EPP) amplitude as the muscle membrane potential (V_m) was altered to the values shown at the left of each record by steady currents injected through a second intracellular microelectrode; calibration pulse, 5 mV, 5 ms. Each point in the graph represents the mean amplitude of 5–10 EPP's. C: ACh response (arrow) evoked by a 1‐ms pulse of ACh precisely mimics a spontaneous synaptic potential recorded on the same trace; calibration, 5 mV, 10 ms. D: reversible reduction in EPP size by d‐tubocurare (10⁻⁷ g/ml); record at extreme right obtained 30 min after removal of the drug; bars, 15 mV, 15 ms. E: miniature end‐plate potentials (MEPP's) recorded on contiguous segments of moving film; bars, 5 mV, 2 s. F: variation in amplitude (and one failure) of successive EPP's in a series evoked at a rate of 1/s, despite a constant stimulus (action potential) in the innervating neuron (upper traces); calibration pulse, 5 mV, 5 ms

A, B, D, and F from Fischbach 91; C from Fischbach & Cohen 93

Figure 23.

A site of transmitter release identified by focal depolarization in the presence of TTX (see text). Interference contrast illumination of a chick muscle cell contacted by a neurite from a nearby spinal cord explant. Synaptic potentials were evoked by extracellular stimulation at α but not at b or elsewhere along the process which extends along the edge of the fiber

From Fischbach et al. 372

Figure 24.

Appearance of ACh receptors in muscle cells. A: increase in α‐bungarotoxin (α‐BGT) binding in control (▪) and in fusion‐arrested (•) chick muscle cultures (see text). ▴, fusion index (percent of nuclei incorporated into myotubes). Δ, fusion index in low‐Ca²⁺ media. Note that the increase in toxin binding lags behind the fusion index in control cultures and is even further delayed when fusion is blocked.

From Patterson & Prives 244.] B: elongated mononucleated muscles from rat muscle grown in standard media for 48 h. The oscilloscope record shows an ACh response (current on lower trace) recorded in cell 1. Horizontal bar, 100 μm. [From Fambrough & Rash 89.] C: autoradiographs of freshly dissociated (without the use of enzymes) mononucleated cells from chick muscle exposed in suspension to [¹²⁵I]α‐BGT. Interference contrast and bright‐field views. Horizontal bar, 10 μm. [From Smilowitz et al. 322

Figure 25.

Synthesis and degradation of ACh receptors by cultured muscle fibers. A: following a saturating exposure to unlabeled α‐bungarotoxin (α‐BGT). The appearance of new receptors was determined by [¹²⁵I]α‐BGT binding (•). ○, cultures treated as above but pretreated for 3 h with cycloheximide; Δ, cultures preincubated in dinitrophenol plus iodoacetate.

From Harzell & Fambrough 128.] B: release of radioactivity into the medium from chick or rat muscle fibers saturated with [¹²⁵I]α‐BGT. Release, expressed as a fraction of that initially bound, is a first‐order process with a half‐time of about 20 h. [From Devreotes & Fambrough 73

Figure 26.

Histograms of ACh sensitivity of uninnervated (A) and innervated (B) muscle fibers in spinal cord muscle cell cultures.

From Fischbach and Cohen 93.] In (C) active and inactive uninnervated fibers are compared. No neurons were present. Active fibers stimulated intermittently for 2–3 days at 10/sec (\ \ \ \) were less sensitive than inactive fibers grown in the presence of TTX (/ / / /). [From Cohen & Fischbach 53. Copyright 1973 by the American Association for the Advancement of Science

Figure 27.

Relative peaks of ACh sensitivity on uninnervated (A) and innervated (B) muscle fiber; ordinate, mV/nC. In A, bars indicate position of hypolemmal muscle nuclei. [From Fischbach & Cohen 93.] B: a hot spot located exactly at a site of transmitter release. Inset, synaptic potentials evoked by focal depolarization at the termination of a nerve process. ACh was iontophoresed at the points indicated in the tracing (prepared from a photograph), and the sensitivities at each point are shown in register below. Bars, 5 mV, 5 ms.

From S. A. Cohen and G. D. Fischbach, unpublished observations

Figure 28.

Tubulovesicular network of cultured muscle fibers. In A note the openings to the exterior and also that some tubules form junctions with cisternae of the sarcoplasmic reticulum (arrow). Horizontal bar, 0.25 μm. B: unstained section of a muscle fiber exposed to horseradish peroxidase prior to fixation. The reaction product outlines areas continuous with the extracellular space. N, nucleus; G, Golgi; T, tubules; S, sarcoplasmic reticulum.

Micrographs provided by M. Henkart

Figure 29.

Spontaneously occurring synaptic potentials in dissociated chick spinal cord cells. A and B: superimposed traces show depolarizing potentials (EPSP's) in one cell (A), hyperpolarizing potentials (IPSP's) in another (B, upper record), and both in a third cell (B, lower record). Vertical bar, 10 mV for A, 12 mV for B. Horizontal bar, 5 ms for A, 10 ms for B, upper trace, 20 ms for B, lower trace. C: a continuous record shows regularly occurring large EPSP's that often trigger action potentials. Note the few small PSP's. D: Small PSP's recorded in the presence of TTX (10⁻⁷ g/ml). Vertical bar, 10 mV for C, 5 mV for D. Horizontal bar, 1 s for C, 5 s for D

From Fischbach & Dichter 95

Figure 30.

Stimulus‐evoked synaptic potentials in mouse spinal cord‐sensory ganglion cell cultures. A: monophasic EPSP (trace 3) evoked in one spinal cord neuron by stimulation (current in trace 1) of another spinal cord neuron (trace 2). B: action potential in a sensory neuron (evoked by a brief current pulse — not shown); trace 1 is followed by a small variable EPSP in a spinal cord cell (superimposed traces in 2). A larger less variable EPSP evoked by another sensory neuron in another spinal cord cell is shown in trace 3. C: complex responses in spinal cord neurons. An outward current plus (trace 3) evokes three spikes and is followed by a long‐lasting barrage of spikes and PSP's in the same (trace 2) and in an adjacent (trace 1) neuron.

Figure 31.

Electron micrographs of synaptic structures in chick spinal cord (A) and rat brain (B and C) cell cultures. In each, note the accumulations of small clear vesicles, the larger dense‐core vesicles, and increased density or thickening of the synaptic membranes. A, Several synaptic profiles appear along a single process. × 18,000. C: a synapse contacts a small process which may be a “spine.’ B and C, × 32,000.

A, from Fischbach & Dichter 95; B and C provided by E. Neale

Figure 32.

Stimulation of a single presynaptic ending on a chick spinal cord neuron. Successive brief negative current pulses, delivered through a 2‐μm‐tipped NaCl‐filled pipette placed exactly over the bouton indicated by the arrow, evoked the synaptic potentials shown at right (traced from photographs). The response disappeared when the stimulating electrode was moved about 5 μm in any direction. Note that the incoming nerve processes pass out of the plane of focus. The postsynaptic nerve cell body overlies a muscle fiber. Calibration bars, 5 mV, 5 ms.

Figure 33.

Drug responses. A: ACh potentials in a sympathetic ganglion cell. The 20‐nA iontophoretic pulse is shown in the second trace; suprathreshold and subthreshold responses are shown in a and b. B: glutamate potential in a spinal cord neuron; iontophoretic pulse, 30 nA. This response is slower than EPSP's recorded in the same cell (trace 3). C: GABA potentials in a sympathetic ganglion cell. D: GABA potentials in a spinal cord neuron. A 25‐nA pulse (duration indicated by bar) was repeated as the membrane potential was shifted. The response reverses in polarity between −70 and −50 mV

A and C from Obata 233; B and D from B. R. Ransom and P. G. Nelson, unpublished observations

Figure 34.

Nonuniform distribution of chemoreceptors on a mouse spinal cord neuron (A) and a neuroblastoma cell (B) about 4 wk after plating. A: penwriter records of glutamate responses at the sites indicated. Note the relatively large and rapid response at lower left. The same neuron impregnated with silver is shown in the inset.

From B. R. Ransom and P. G. Nelson, unpublished observations.) B: ACh produces a qualitatively different response at different sites — a depolarization over the soma, a hyperpolarization at relatively distant sites along processes, and a biphasic response at intermediate points. (From Peacock et al. 1973

Figure 35.

A: increase in synthesis of catecholamines (norepinephrine and dopamine) from [³H]tyrosine by rat sympathetic ganglion neurons with time in culture. Note the initial log of 7–8 days followed by a 50‐fold increase. This pattern is markedly different from the increase in total accumulation of radioactivity measured in the same cultures (B)

From Mains & Patterson 196

Figure 36.

Cholinergic synapses between sympathetic ganglion cells. A: simultaneous recordings from two sympathetic neurons in which each spike in one (the “driver,’ middle trace) is followed by a depolarizing synaptic potential in the other (the “follower,’ upper trace). Some PSP's in the follower are large enough to trigger spikes. The first spike in the driver cell is evoked by a brief pulse of outward current (lower trace). The other spikes arise spontaneously. B: the same cells in the presence of 50 μM hexamethonium, a nicotinic cholinergic antagonist. All spontaneous PSP's are abolished, and the evoked spike in the driver neuron is no longer followed by a PSP in the follower. C: the same cells after washing out the hexamethonium. Calibration bars: 40 mV, 40 ms.

From O. MacLeish, P. H. O'Lague, E. J. Furshpan, and D. D. Potter, unpublished observations

References
1.	Abercrombie, M., J. E. Heaysman, and S. M. Pegrun. The locomotion of fibroblasts in culture. I. Movements of the leading edge. Exptl. Cell Res. 59: 393–398, 1970.
2.	Abercrombie, M., J. E. Heaysman, and S. M. Pegrun. The locomotion of fibroblasts in culture. II. “Ruffling.” Exptl. Cell Res. 60: 437–444, 1970.
3.	Albers, R. W., G. J. Siegel, R. Katzmann, and B. W. Agranoff (editors)., Basic Neurochemistry. Boston: Little, Brown, 1972.
4.	Amano, T., E. Richelson, and M. Nirenberg. Neurotransmitter synthesis by neuroblastoma clones. Proc. Natl. Acad. Sci. US 69: 258–263, 1972.
5.	Amsterdam, A., and J. D. Jamieson. Studies on dispersed pancreatic exocrine cells. I. Dissociation technique and morphologic characteristics of separated cells. J. Cell Biol. 63: 1037–1056, 1974.
6.	Amsterdam, A., and J. D. Jamieson. Studies on dispersed pancreatic exocrine cells. II. Functional characteristics of separated cells. J. Cell Biol. 63: 1057–1073, 1974.
7.	Anderson, C. R., and C. F. Stevens. Voltage clamp analysis of acetylcholine produced end‐plate current fluctuations at frog neuromuscular junctions. J. Physiol. London 235: 655–691, 1973.
8.	Augusti‐Tocco, G., and G. Sato. Establishment of functional clonal lines of neurons from mouse neuroblastoma. Proc. Natl. Acad. Sci. US 64: 311–315, 1969.
9.	Balinsky, B. L. Introduction to Embryology (4th ed.). Philadelphia: Saunders, 1975.
10.	Balsamo, J., and J. Lilien. Embryonic cell aggregation: kinetics and specificity of binding of enhancing factors. Proc. Natl. Acad. Sci. US 71: 727–731, 1974.
11.	Balsamo, J., and J. Lilien. Functional identification of three components which mediate tissue type specific embryonic cell adhesion. Nature 251: 522–524, 1974.
12.	Balsamo, J., J. McDonough, and J. Lilien. Retinal tectal connections in embryonic chick: evidence for regionally specific cell surface components which mimic the pattern of innervation. Devel. Biol. 49: 338–346, 1976.
13.	Barbera, A. J., R. B. Marchase, and S. Roth. Adhesive recognition and retinotectal specificity. Proc. Natl. Acad. Sci. US 70: 2482–2486, 1973.
14.	Barker, J. L., and R. A. Nicoll. GABA: role in primary afferent depolarization. Science 176: 1043–1045, 1972.
15.	Barkley, D. S., L. L. Rakie, J. K. Chaffee, and D. L. Wong. Cell separation by velocity sedimentation of postnatal mouse cerebellum. J. Cellular Physiol. 81: 271–280, 1973.
16.	Barnard, E. A., J. Wieckowsk, and T. H. Chiu. Cholinergic receptor molecules and cholinesterase molecules at mouse skeletal muscle junctions. Nature 234: 207–209, 1971.
17.	Behnke, O., B. Kristensen, and L. Nielsen. Electron microscopical observations on actinoid and myosinoid filaments in blood platelets. J. Ultrastruct. Res. 37: 351–369, 1971.
18.	Benda, P. J., J. Lightbody, G. H. Sato, L. Levine, and W. Sweet. Differentiated rat glial strain in tissue culture. Science 161: 370–371, 1968.
19.	Bennett, M. R., E. M. McLachlan, and R. S. Taylor. The formation of synapses in reinnervated mammalian striated muscle. J. Physiol. London 233: 481–500, 1973.
20.	Bennett, M. R., and A. G. Pettigrew. The formation of synapses in striated muscle during development. J. Physiol. London 241: 515–545, 1974.
21.	Bennett, M. V. L. Function of electrotonic junctions in embryonic and adult tissues. Federation Proc. 32: 65–75, 1973.
22.	Berg, D. K., and G. D. Fischbach. Spinal cord cells: attempts at fractionation (Abstract). Federation Proc. 34: 405, 1975.
23.	Berg, D. K., and Z. W. Hall. Fate of α‐bungarotoxin bound to acetylcholine receptors of normal and denervated muscle. Science 184: 473–474, 1974.
24.	Berry, M. N., and D. S. Friend. High‐yield preparation of isolated rat liver parenchymal cells. J. Cell Biol. 43: 506–520, 1969.
25.	Betz, W., and B. Sakmann. Effects of proteolytic enzymes in function and structure of frog neuromuscular junctions. J. Physiol. London 230: 673–688, 1973.
26.	Bischoff, R., and H. Holtzer. Inhibition of myoblast fusion after one round of DNA synthesis in 5‐bromodeoxyuridine. J. Cell Biol. 44: 134–150, 1970.
27.	Blume, A., F. Gilbert, S. Wilson, J. Farber, R. Rosenberg, and M. Nirenberg. Regulation of acetylcholinesterase in neuroblastoma cells. Proc. Natl. Acad. Sci. US 67: 786–792, 1970.
28.	Bodian, D. Development of fine structure of spinal cord in monkey fetuses. I. The motoneuron neuropil at the time of onset of reflex activity. Bull. Johns Hopkins Hosp. 119: 129–149, 1966.
29.	Boethius, J., and E. Knuttson. Resting membrane potential in chick muscles during ontogeny. J. Exptl. Zool. 174: 281–286, 1970.
30.	Bolton, T. B. The role of electrogenic sodium pumping in the response of smooth muscle to acetylcholine. J. Physiol. London 228: 713–731, 1973.
31.	Bornstein, M. B. Reconstituted rat‐tail collagen as a substrate for tissue cultures on coverslips in Maximow slides and in roller tubes. Lab. Invest. 7: 134–137, 1958.
32.	Bray, D. Surface movements during the growth of single explanted neurons. Proc. Natl. Acad. Sci. US 65: 905–910, 1970.
33.	Bray, D. Branching patterns of individual sympathetic neurons in culture. J. Cell Biol. 56: 702–712, 1973.
34.	Bray, D. Model for membrane movements in the neural growth cone. Nature 244: 93–95, 1973.
35.	Breakefield, X. O., and M. W. Nirenberg. Selection for neuroblastoma clones that synthesize certain transmitters. Proc. Natl. Acad. Sci. US 71: 2530–2533, 1974.
36.	Buckley, P. A., and I. R. Konigsberg. Myogenic fusion and the duration of the post‐mitotic gap (G1). Develop. Biol. 37: 193–212, 1974.
37.	Bunge, M. B. Fine structure of nerve fibers and growth cones of isolated sympathetic neurons in culture. J. Cell Biol. 56: 713–735, 1973.
38.	Bunge, M. B., R. P. Bunge, and E. R. Peterson. The onset of synapse formation in spinal cord cultures as studied by electron microscopy. Brain Res. 6: 728–749, 1967.
39.	Bunge, R. P., R. Rees, P. Wood, H. Burton, and C. P. Ko. Anatomical and physiological observations on synapses formed on isolated autonomic neurons in tissue culture. Brain Res. 66: 401–412, 1974.
40.	Burke, R., and P. G. Nelson. Synaptic activity in motoneurons during natural stimulation of muscle spindles. Science 151: 1088–1091, 1966.
41.	Catterall, W. A., and M. Nirenberg. Sodium uptake associated with activation of action potential ionophores of cultured neuroblastoma and muscle cells. Proc. Natl. Acad. Sci. US 70: 3759–3763, 1973.
42.	Chalazonitis, A., and L. A. Greene. Enhancement in excitability properties of mouse neuroblastoma cells cultured in the presence of dibutyryl cyclic AMP. Brain Res. 72: 340–345, 1974.
43.	Chalazonitis, A., L. A. Greene, and M. Nirenberg. Electrophysiological characteristics of chick embryo sympathetic neurons in dissociated cell culture. Brain Res. 68: 235–252, 1974.
44.	Chalazonitis, A., L. Greene, and W. Shain. Excitability and chemosensitivity properties of a somatic cell hybrid between mouse neuroblastoma and sympathetic ganglion cells. Exptl. Cell Res. 96: 225–238, 1975.
45.	Chamley, J. H., G. R. Campbell, and G. Burnstock. An analysis of the interactions between sympathetic nerve fibers and smooth muscle cells in tissue culture. Develop. Biol. 33: 344–361, 1973.
46.	Chamley, J. H., I. Goller, and G. Burnstock. Selective growth of sympathetic nerve fibers to explants of normally densely innervated autonomic effector organs in tissue culture. Develop. Biol. 31: 362–379, 1973.
47.	Chang, S.‐Y., and C. Kung. Temperature‐sensitive pawns: conditional behavioral mutants of Paramecium aurelia. Science 180: 1197–1199, 1973.
48.	Chen, J. S., and R. Levi‐Montalcini. Axonal outgrowth and cell migration in vitro from nervous system of cockroach embryos. Science 166: 631–632, 1969.
49.	Chen, J. S., and R. Levi‐Montalcini. Long‐term cultures of dissociated nerve cells from the embryonic nervous system of the cockroach Periplaneta americana. Arch. Ital. Biol. 108: 503–537, 1970.
50.	Claude, P., and J. G. Hildebrand. Lack of accumulation of transmitter compounds by neuroblastoma cells in monolayer culture. J. Cell Biol. 55: 102a, 1971.
51.	Cohen, M. W. The development of neuromuscular connections in the presence of d‐tubocurare. Brain Res. 41: 457–463, 1972.
52.	Cohen, M. W., and M. J. Anderson. Fluorescent staining of acetylcholine receptors in vertebrate skeletal muscle. J. Physiol. London 237: 385–400, 1973.
53.	Cohen, S. A., and G. D. Fischbach. Regulation of muscle acetylcholine sensitivity by muscle activity in cell culture. Science 181: 76–78, 1973.
54.	Cohen, S. A., and G. D. Fischbach. ACh sensitivity at nerve‐muscle synapses in vitro (Abstract). Federation Proc. 34: 405, 1975.
55.	Coleman, J., and A. Coleman. Muscle differentiation and macromolecular synthesis. J. Cellular Physiol. 72, Suppl. 1: 19–34, 1968.
56.	Cone, C. D. Unified theory on the basic mechanism of normal mitotic central oncogenesis. J. Theoret. Biol. 30: 151–181, 1971.
57.	Cone, C. D., and M. Tangier, Jr. Contact inhibition of division: involvement of the electrical transmembrane potential. J. Cellular Physiol. 82: 373–386, 1973.
58.	Coon, H. G. Clonal stability and phenotypic expression of chick cartilage cells in vitro. Proc. Natl. Acad. Sci. US 55: 66–73, 1966.
59.	Crain, S. M. Resting and action potentials of cultured chick embryo spinal ganglion cells. J. Comp. Neurol. 104: 285–330, 1956.
60.	Crain, S. M. Development of “organotypic” bioelectric activities in central nervous tissue during maturation in culture. Intern. Rev. Neurobiol. 9: 1–43, 1966.
61.	Crain, S. Intracellular recordings suggesting synaptic functions in chick embryo spinal sensory ganglion cells isolated in vitro. Brain Res. 26: 188–191, 1971.
62.	Crain, S., M. B. Bornstein, and E. R. Peterson. Maturation of cultured embryonic CNS tissues during chronic exposure to agents which prevent bioelectric activity. Brain Res. 8: 363–372, 1968.
63.	Crain, S. M., and E. R. Peterson. Development of paired explants of fetal spinal cord and adult skeletal muscle during chronic exposure to curare and hemicholinium. In Vitro 6: 373, 1971.
64.	Daniels, M. P. Colchicine inhibition of nerve fiber formation in vitro. J. Cell Biol. 53: 164–176, 1972.
65.	Daniels, M., and S. Vogel. Immunoperoxidase staining of α‐bungarotoxin binding sites in muscle endplates shows distribution of acetylcholine receptors. Nature 254: 339–341, 1975.
66.	Davidson, R. L. Regulation of gene expression in somatic cell hybrids: a review. In Vitro 6: 411–426, 1971.
67.	De Groat, W. C. GABA‐depolarization of a sensory ganglion: antagonism by picrotoxin and bicuculline. Brain Res. 38: 429–432, 1972.
68.	del Castillo, J., and B. Katz. Localization of active spots within the neuromuscular junction of the frog. J. Physiol. London 132: 630–649, 1956.
69.	del Cerro, M. P., and R. S. Snider. Studies on the developing cerebellum. Ultrastructure of the growth cones. J. Comp. Neurol. 133: 341–362, 1968.
70.	Dennis, M. J. Physiologic properties of junctions between nerve and muscle developing during salamander limb regeneration. J. Physiol. London 244: 683–702, 1975.
71.	Dennis, M. J., and R. Miledi. Characteristics of transmitter release at regenerating frog neuromuscular junctions. J. Physiol. London 239: 571–594, 1974.
72.	Dennis, M. J., and R. Miledi. Non‐transmitting neuromuscular junctions during an early stage of end‐plate reinnervation. J. Physiol. London 239: 553–570, 1974.
73.	Devreotes, P. N., and D. M. Fambrough. Acetylcholine receptor turnover in membranes of developing muscle fibers. J. Cell Biol. 65: 335–358, 1975.
74.	Diamond, J., and R. Miledi. A study of fetal and newborn rat muscle fibers. J. Physiol. London 162: 373–408, 1962.
75.	Dichter, M. A., and G. D. Fischbach. Functional synapses in cell cultures of chick spinal cord and dorsal root ganglia. Abstr. Ann. Meeting Am. Soc. Cell Biol., 4th, 1971, p. 76a.
76.	Dichter, M., and G. D. Fischbach. The action potential of chick dorsal root ganglion neurons maintained in cell culture. J. Physiol. London. In press.
77.	Druckrey, H., S. Ivankovic, R. Preussmann, K. J. Zulch, and H. D. Mennel. Selective induction of malignant tumors of the nervous system by resorptive carcinogens. In: Experimental Biology of Brain Tumors, edited by W. M. Kirsch. Springfield, Ill.: Thomas, 1972, p. 85–147.
78.	Eagle, H. The specific amino acid requirements of a mammalian cell (strain L) in tissue culture. J. Biol. Chem. 214: 839–852, 1955.
79.	Eagle, H., and K. A. Piez. The population‐dependent requirement by cultured mammalian cells for metabolites which they can synthesize. J. Exptl. Med. 116: 29–43, 1962.
80.	Ebert, J., and I. M. Sussex. Interacting Systems in Development (2nd ed.). New York: Holt, Reinhart and Winston, 1970.
81.	Eccles, J. C. The Physiology of Synapses. New York: Academic, 1964.
82.	Eckert, R., and Y. Naitoh. Bioelectric control of locomotion in the ciliates. J. Protozool. 19: 237–243, 1972.
83.	Ehrmann, R. L., and G. O. Gey. The growth of cells on a transparent gel of reconstituted rat‐tail collagen. J. Natl. Cancer Inst. 16: 1375–1404, 1956.
84.	Elsdale, T., and J. Bard. Collagen substrata for studies on cell behavior. J. Cell Biol. 54: 626–637, 1972.
85.	Emerson, C. P., Jr., and S. L. Beckner. Activation of myosin synthesis in fusing and mononucleated myoblasts. J. Mol. Biol. 93: 431–448, 1975.
86.	Ephrussi, B. Hybridization of Somatic Cells. Princeton: Princeton Univ. Press, 1972.
87.	Ezerman, E. B., and H. Ishikawa. Differentiation of the sarcoplasmic reticulum and T system in developing chick skeletal muscle in vitro. J. Cell Biol. 35: 405–420, 1967.
88.	Fambrough, D. M., Cellular and developmental biology of acetylcholine receptors in skeletal muscle. In: Neurochemistry of Cholinergic Receptors, edited by E. De Robertis and J. Schacht. New York: Raven, 1974.
89.	Fambrough, D. M., H. C. Hartzell, J. A. Powell, J. E. Rash, and N. Joseph. On the differentiation and organization of the surface membrane of a post‐synaptic cell — the skeletal muscle fiber. In: Synaptic Transmission and Neuronal Interaction, edited by M. V. L. Bennett. New York: Raven, 1974.
90.	Fambrough, D., H. C. Hartzell, J. E. Rash, and A. K. Ritchie. Receptor properties of developing muscle. Ann. N.Y. Acad. Sci. 228: 47–62, 1974.
91.	Fambrough, D., and J. E. Rash. Development of acetylcholine sensitivity during myogenesis. Develop. Biol. 26: 55–68, 1971.
92.	Fischbach, G. D. Synaptic potentials recorded in cell cultures of nerve and muscle. Science 169: 1331–1333, 1970.
93.	Fischbach, G. D. Synapse formation between dissociated nerve and muscle cells in low density cultures. Develop. Biol. 28: 407–429, 1972.
94.	Fischbach, G. D., Some aspects of neuromuscular junction formation. In: Cell Communication, edited by R. P. Cox. New York: Wiley, 1974, p. 43–66.
95.	Fischbach, G. D., D. K. Berg, S. A. Cohen, and E. Frank. Enrichment of nerve‐muscle synapses in spinal cord‐muscle cultures and identification of relative peaks of ACh sensitivity at sites of transmitter release. Cold Spring Harbor Symp. Quant. Biol. 40: 347–357, 1976.
96.	Fischbach, G. D., and S. A. Cohen. The distribution of acetylcholine sensitivity over uninnervated and innervated muscle fibers grown in cell culture. Develop. Biol. 31: 147–162, 1973.
97.	Fischbach, G. D., S. A. Cohen, and M. P. Henkart. Some observations on trophic interaction between neurons and muscle fibers in cell culture. Ann. NY Acad. Sci. 228: 35–46, 1974.
98.	Fischbach, G. D., and M. A. Dichter. Electrophysiologic and morphologic properties of neurons in dissociated chick spinal cord cell cultures. Develop. Biol. 37: 100–116, 1974.
99.	Fischbach, G. D., D. Fambrough, and P. G. Nelson. A discussion of neuron and muscle cells in culture. Federation Proc. 32: 1636–1642, 1973.
100.	Fischbach, G. D., M. P. Henkart, S. A. Cohen, A. C. Breuer, J. A. Whysner, and F. M. Neal. Studies on the development of neuromuscular junctions in cell culture. In: Synaptic Transmission and Neuronal Interaction, edited by M. V. L. Bennett. New York: Raven, 1974, p. 259–285.
101.	Fischbach, G. D., M. Nameroff, and P. G. Nelson. Electrical properties of chick skeletal muscle fibers developing in cell culture. J. Cellular Physiol. 68: 289–300, 1971.
102.	Fischman, D. A., Development of striated muscle. In: The Structure and Function of Muscle (2nd ed.), edited by G. H. Bourne. New York: Academic, 1972, vol. I, p. 75–149.
103.	Fluck, R. A., and R. C. Strohman. Acetylcholinesterase activity in developing skeletal muscle cells in vitro. Develop. Biol. 33: 417–428, 1973.
104.	Frazier, W. A., C. E. Ohlendorf, L. F. Boyd, L. Aloe, E. M. Johnson, J. A. Ferrendelli, and R. A. Bradshaw. Mechanism of actions of nerve growth factor and cyclic AMP on neurite outgrowth in embryonic chick sensory ganglia: demonstration of independent pathways of stimulation. Proc. Natl. Acad. Sci. US 70: 2448–2452, 1973.
105.	Fukuda, J. Chloride spike: a third type of action potential in tissue‐cultured skeletal muscle cells from the chick. Science 185: 76–78, 1974.
106.	Furmanski, P., D. J. Silverman, and M. Lobin. Expression of differentiated functions in mouse neuroblastoma mediated by dibutyryl cyclic AMP. Nature 233: 413–415, 1971.
107.	Furshpan, E. J., and D. D. Potter. Low‐resistance junctions between cells in embryos and tissue culture. In: On Current Topics in Developmental Biology, edited by A. A. Moscona and A. Monroy. New York: Academic, 1968, vol. 3, p. 95–127.
108.	Garber, B., and A. A. Moscona. Reconstruction of brain tissue from cell suspensions. I. Aggregation patterns of cells dissociated from different regions of the developing brain. Develop. Biol. 27: 217–234, 1972.
109.	Giller, E. L., X. O. Breakefield, C. N. Christian, E. A. Neale, and P. G. Nelson. Expression of neuronal characteristics in culture: some pros and cons of primary cultures and continuous cell lines. In: Perspectives in Neurobiology, edited by M. Santini. New York: Raven, 1975, p. 603–623.
110.	Giller, E. L., Jr., B. K. Schrier, A. Shainberg, H. R. Fisk, and P. G. Nelson. Choline acetyltransferase activity is increased in combined cultures of spinal cord and muscle cells from mice. Science 182: 588–589, 1973.
111.	Glick, M. C., Y. Kimhi, and U. Z. Littauer. Glycopeptides from surface membranes of neuroblastoma cells. Proc. Natl. Acad. Sci. US 70: 1682–1687, 1973.
112.	Godfrey, E. W., P. G. Nelson, B. K. Schrier, A. C. Breuer, and B. R. Ransom. Neurons from fetal rat brain in a new cell culture system: a multidisciplinary analysis. Brain Res. 90: 1–21, 1975.
113.	Goldstein, M. N., and D. Pinkel. Long‐term tissue culture of neuroblastomas. J. Natl. Cancer Inst. 20: 675–689, 1958.
114.	Gorman, A. L. F., and M. F. Marmor. Steady state contribution of the sodium pump to the resting potential of molluscan neurone. J. Physiol. London 242: 35–48, 1974.
115.	Gottlieb, D. I., R. Merrell, and L. Glazer. Temporal changes in embryonic cell surface recognition. Proc. Natl. Acad. Sci. US 21: 1800–1802, 1974.
116.	Green, M., and H. Neurath. The effects of divalent cations on trypsin. J. Biol. Chem. 204: 379, 1953.
117.	Greene, L. A., A. J. Sytkowski, Z. Vogel, and M. Nirenberg. α‐Bungarotoxin used as a probe for acetylcholine receptors of cultured neurons. Nature 243: 163–166, 1973.
118.	Greene, L. A., W. Shain, A. Chalazonitis, X. Breakfield, J. Minna, H. Coon, and M. W. Nirenberg. Neuronal properties of hybrid neuroblastoma X sympathetic ganglion cells. Proc. Natl. Acad. Sci. US 72: 4923–4927, 1975.
119.	Guillery, R. W., H. M. Sobkowicz, and G. L. Scott. Relationships between glial and neuronal elements in the development of long‐term cultures of the spinal cord of the fetal mouse. J. Comp. Neurol. 140: 1–34, 1970.
120.	Gutmann, E., and J. Z. Young. The re‐innervation of muscle after various periods of atrophy. J. Anat. 78: 15–43, 1944.
121.	Hall, Z., and R. B. Kelly. Enzymatic detachment of end‐plate acetylcholinesterase from muscle. Nature New Biol. 232: 62–63, 1971.
122.	Ham, R. G., Cloning of mammalian cells. In: Methods in Cell Physiology, edited by D. M. Prescott. New York: Academic, 1972, vol. V, 37–74.
123.	Ham, R. G. Nutritional requirements of primary cultures. A neglected problem of modern biology. In Vitro 10: 119–129, 1974.
124.	Hamburger, V. Origins of integrated behavior. Develop. Biol. Suppl. 2: 251–271, 1968.
125.	Harris, A. J. Inductive functions of the nervous system. Ann. Rev. Physiol. 36: 251–305, 1974.
126.	Harris, A. J., Role of acetylcholine receptors in synapse formation. In: Synaptic Transmission and Neuronal Interaction, edited by M. V. L. Bennett. New York: Raven, 1974, p. 315–337.
127.	Harris, A. J., and M. J. Dennis. Acetylcholine sensitivity and distribution on mouse neuroblastoma. Science 167: 1253–1255, 1970.
128.	Harris, A. J., S. Heinemann, D. Schubert, and H. Tarakis. Trophic interaction between cloned tissue culture lines of nerve and muscle. Nature 231: 296–301, 1971.
129.	Harris, A. J., S. W. Kuffler, and M. J. Dennis. Differential chemosensitivity of synaptic and extrasynaptic areas on the neuronal surface membrane of parasympathetic neurons of the frog, tested by microapplication of acetylcholine. Proc. Roy. Soc. London Ser. B 177: 541–553, 1971.
130.	Harris, A. K., Cell surface movements related to locomotion. In: Locomotion of the Tissue Cells. New York: Elsevier, 1973, p. 3–26. (Ciba Foundation Symp. 14.)
131.	Harris, J. B., and M. W. Marshall. Tetrodotoxin resistant action potentials in newborn rat muscle. Nature New Biol. 243: 191–192, 1973.
132.	Harrison, R. G. The outgrowth of the nerve fiber as a mode of protoplasmic movement. J. Exptl. Zool. 9: 787–846, 1910.
133.	Hartzell, H. C., and D. M. Fambrough. Acetylcholine receptor production and incorporation into membranes of developing muscle fibers. Develop. Biol. 30: 153–165, 1973.
134.	Hauschka, S. D., and I. Konigsberg. The influence of collagen on the development of muscle clones. Proc. Natl. Acad. Sci. US 55: 119–126, 1966.
135.	Hazelwood, C. F., and B. L. Nichols. The in vivo muscle resting potential in the developing rat. Johns Hopkins Med. J. 123: 198–203, 1968.
136.	Hazelwood, C. F., and B. L. Nichols. Changes in muscle sodium, potassium chloride, water and voltage during maturation in the rat: an experimental and theoretical study. Johns Hopkins Med. J. 125: 119–132, 1969.
137.	Heilbrunn, L. V. An Outline of General Physiology (2nd ed.). Philadelphia: Saunders, 1943.
138.	Henn, F. A., and A. Hamberger. Glial cell function; uptake of transmitter substances. Proc. Natl. Acad. Sci. US 68: 2686–2690, 1971.
139.	Herrup, K., and E. M. Shooter. Properties of the B nerve growth factor receptor of avian dorsal root ganglia. Proc. Natl. Acad. Sci. US 70: 3884–3888, 1973.
140.	Herschman, H. R., B. P. Grauling, and M. P. Lerner. Nervous system specific proteins in cultured neural cells. In: Tissue Culture of the Nervous System, edited by G. Sato. New York: Plenum, 1973, p. 187–202.
141.	Hier, D. B., B. G. W. Arnason, and M. Young. Studies on the mechanism of action of nerve growth factor. Proc. Natl. Acad. Sci. US 69: 2268–2272, 1972.
142.	Hild, W., and I. Tasaki. Morphological and physiological properties of neurons and glial cells in tissue culture. J. Neurophysiol. 25: 277–304, 1962.
143.	Hodgkin, A. L., and B. Katz. The effect of sodium ions on the electrical activity of the giant axon of the squid. J. Physiol. London 108: 37–39, 1949.
144.	Holtzer, H., Myogenesis. In: Cell Differentiation, edited by O. Scheide and J. de Veillis. Princeton: Van Nostrand, 1970, p. 476–503.
145.	Hopkins, C. R., and M. G. Farquhar. Hormone secretion by cells dissociated from rat anterior pituitaries. J. Cell Biol. 59: 276–303, 1973.
146.	Hourani, B. T., B. F. Torain, M. P. Henkart, R. L. Carter, V. T. Marchesi, and G. D. Fischbach. Acetylcholine receptors of cultured muscle cells demonstrated with ferritin‐α‐bungarotoxin conjugates. J. Cell Sci. 16: 473–479, 1974.
147.	Howard, R. B., J. C. Lee, and L. A. Pesch. The fine structure, potassium content and respiratory activity of isolated rat liver parenchyma cells prepared by improved enzymatic techniques. J. Cell Biol. 57: 642–658, 1973.
148.	Hughes, A. The growth of embryonic neurites — a study on cultures of chick neural tissue. J. Anat. 87: 150–162, 1953.
149.	Huxley, H. Muscular contraction and cell motility. Nature 243: 445–449, 1973.
150.	Ishikawa, H. Formation of elaborate networks of T‐system tubules in cultured skeletal muscle with special reference to the T‐system formation. J. Cell Biol. 38: 51–66, 1968.
151.	Ito, M. The electrical activity of spinal ganglion cells investigated with intracellular microelectrodes. Japan. J. Physiol. 7: 297–323, 1957.
152.	Jacobson, M. Developmental Neurobiology. New York: Holt, Reinhart, and Winston, 1970.
153.	James, D. W., and R. L. Tresman. An electron microscopic study of the de novo formation of neuromuscular junctions in tissue culture. Z. Zellforsch. Mikroskop. Anat. 106: 126–140, 1969.
154.	Jankowska, E., and S. Lindstrom. Morphologic identification of physiologically defined neurones in the cat spinal cord. Brain Res. 20: 323–326, 1970.
155.	Johnson, D. G., P. Gorden, and I. J. Kopin. A sensitive radioimmunoassay for 75 nerve growth factor antigens in serum and tissues. J. Neurochem. 18: 2355–2362, 1971.
156.	Johnson, D. G., S. D. Silberstein, I. Hanbauer, and I. J. Kopin. The role of nerve growth factor in the ramification of sympathetic nerve fibers into the rat iris in organ culture. J. Neurochem. 19: 2025–2029, 1972.
157.	Junge, D., and J. Miller. Different spike mechanisms in axon and soma of molluscan neuron. Nature 252: 155–156, 1974.
158.	Kano, M., and Y. Shimada. Innervation and acetylcholine sensitivity of skeletal muscle differentiated in vitro from chick embryo. J. Cellular Physiol. 78: 233–242, 1971.
159.	Kano, M., and Y. Shimada. Innervation of skeletal muscle cells differentiated in vitro from chick embryo. Brain Res. 27: 402–405, 1971.
160.	Kano, M., and Y. Shimada. Tetrodotoxin‐resistant electric activity in chick skeletal muscle cells differentiated in vitro. J. Cellular Physiol. 81: 85–90, 1972.
161.	Kasa, P., S. P. Mann, and C. Hebb. Localization of choline acetyltransferase. Nature 226: 812–816, 1971.
162.	Kater, S. B., and C. Nicholson. Intracellular Staining in Neurobiology. New York: Springer Verlag, 1973.
163.	Katz, B. Nerve, Muscle and Synapses. New York: McGraw‐Hill, 1966.
164.	Katz, B., and R. Miledi. A study of synaptic transmission in the absence of nerve impulses. J. Physiol. London 192: 407–436, 1967.
165.	Katz, B., and R. Miledi. Further study of the role of calcium in synaptic transmission. J. Physiol. London 207: 789–801, 1970.
166.	Katz, B., and R. Miledi. The statistical nature of the acetylcholine potential and its molecular components. J. Physiol. London 224: 665–699, 1972.
167.	Kawana, E., C. Sandrs, and K. Akert. Ultrastructure of growth cones in the cerebellar cortex of the neonatal rat and cat. Z. Zellforsch. Mikroskop. Anat. 115: 284–298, 1971.
168.	Kelly, A. M., and S. J. Zacks. The fine structure of motor endplate morphogenesis. J. Cell Biol. 42: 154–169, 1969.
169.	Kelly, J. P., and D. C. Van Essen. Cell structure and function in the visual cortex of the cat. J. Physiol. London 238: 515–548, 1974.
170.	Kidokoro, Y. Development of action potentials in a clonal rat skeletal muscle cell line. Nature New Biol. 241: 158–159, 1973.
171.	Kidokoro, Y. Developmental changes of membrane electrical properties in a rat skeletal muscle cell line. J. Physiol. London 244: 129–144, 1975.
172.	Kidokoro, Y. Sodium and calcium components of the action potential in a developing skeletal muscle line. J. Physiol. London 244: 145–160, 1975.
173.	Kidokoro, Y., and S. Heinemann. Synapse formation between clonal muscle cells and rat spinal cord explants. Nature 252: 593–594, 1974.
174.	Klebe, R. J., and F. H. Ruddle. Neuroblastoma: cell culture analysis of a differentiating stem cell system. J. Cell Biol. 43: 69A, 1969.
175.	Koketsu, K., and S. Nishi. Calcium and action potentials of bullfrog sympathetic ganglion cells. J. Gen. Physiol. 53: 608–623, 1969.
176.	Konigsberg, I. R. Clonal analysis of myogenesis. Science 140: 1273–1284, 1963.
177.	Konigsberg, I. R., The relationship of collagen to the clonal development of embryonic skeletal muscle. In: Chemistry and Molecular Biology of the Intercellular Matrix, edited by E. A. Balazs. New York: Academic, 1970, vol. 3, p. 1779–1810.
178.	Kruse, P. F., and M. K. Patterson. Tissue Culture: Methods and Applications. New York: Academic, 1973.
179.	Kuffler, S. W., M. J. Dennis, and A. J. Harris. The development of chemosensitivity in extrasynaptic areas of the neuronal surface after denervation of parasympathetic ganglion cells in the heart of the frog. Proc. Roy. Soc. London Ser. B 177: 555–563, 1971.
180.	Kuffler, S. W., and J. G. Nicholls. The physiology of neuroglial cells. Ergeb. Physiol. Biol. Chem. Exptl. Pharmakol. 57: 1–90, 1966.
181.	Kullberg, R. W. Onset and development of synaptic activity at an amphibian neuromuscular junction (Ph.D. thesis). Montreal, Quebec: McGill University, 1974.
182.	Kung, C. Genetic mutants with altered system of excitation in Paramecium aurelia. I. Phenotypes of the behavioural mutant. Z. Vergleich. Physiol. 7: 142–164, 1971.
183.	Kung, C., and R. Eckert. Genetic modification of electrical properties in an excitable membrane. Proc. Natl. Acad. Sci. US 69: 93–97, 1972.
184.	Kuno, M. Quantum aspects of central and ganglionic synaptic transmission in vertebrates. Physiol. Rev. 51: 647–678, 1971.
185.	Kuno, M., S. Turkanis, and J. Weakly. Correlation between nerve terminal size and transmitter release at the neuromuscular junction of the frog. J. Physiol. London 213: 545–556, 1971.
186.	Land, B. R., P. Raudin, M. Salpeter, and T. R. Podleski. Correlation between acetylcholine sensitivity and density of receptor sites. J. Cell Biol. 63: 183a, 1974.
187.	Lasher, R. S. The uptake of [3H]GABA and differentiation of stellate neurons in cultures of dissociated postnatal rat cerebellum. Brain Res. 69: 235–254, 1974.
188.	Lasher, R. S., and I. S. Zagon. The effect of potassium on neuronal differentiation in cultures of dissociated newborn rat cerebellum. Brain Res. 41: 482–488, 1972.
189.	La Vail, J. H., and M. M. La Vail. Retrograde axonal transport in the central nervous system. Science 176: 1416–1417, 1972.
190.	Letinsky, M. S. The development of nerve‐muscle junctions in Rana catesbiana tadpoles. Develop. Biol. 40: 129–153, 1974.
191.	Levi‐Montalcini, R., and P. U. Angeletti. Nerve growth factor. Physiol. Rev. 48: 534–569, 1968.
192.	Levi‐Montalcini, R., and K. A. Seshan. Long‐term cultures of embryonic and mature insect nervous and neuroendocrine systems. In: Tissue Culture of the Nervous System, edited by G. Sato. New York: Plenum, 1973, p. 1–34.
193.	Lewis, W. H., and M. R. Lewis. The cultivation of sympathetic nerves from the intestine of chick‐embryos in saline solutions. Anat. Record 6: 7–32, 1912.
194.	Lightbody, L., S. Pfeiffer, P. Kornblith, and H. Herschman. Biochemically differentiated clonal glial cells in tissue culture. J. Neurobiol. 1: 411–417, 1970.
195.	Lømø, T., and J. Rosenthal. Control of ACh sensitivity by muscle activity in the rat. J. Physiol. London 221: 493–513, 1972.
196.	Longo, A. M., and E. E. Penhoet. Nerve growth factor in rat glioma cells. Proc. Natl. Acad. Sci. US 71: 2347–2349, 1974.
197.	Luduena, M. A. The growth of spinal ganglion neurons in serum‐free medium. Develop. Biol. 33: 470–476, 1973.
198.	Luduena, M. A. Nerve cell differentiation in vitro. Develop. Biol. 33: 268–284, 1973.
199.	Luduena, M. A., and N. K. Wessels. Cell locomotion, nerve elongation and microfilaments. Develop. Biol. 30: 427–440, 1973.
200.	Macagno, E. R., V. Lopresti, and C. Levinthal. Structure and development of neuronal connections in isogenic organisms: variations and similarities in the optic system of Daphnia magna. Proc. Natl. Acad. Sci. US 10: 57–61, 1973.
201.	Mains, R. Tissue culture of dissociated primary rat sympathetic neurons: studies of growth neurotransmitter metabolism and maturation (Ph.D. thesis). Cambridge: Harvard University, 1973.
202.	Mains, R. E., and P. H. Patterson. Primary cultures of dissociated sympathetic neurons. I. Establishment of long‐term growth in cultures and studies of differentiated properties. J. Cell Biol. 59: 329–345, 1973.
203.	Mains, R. E., and P. H. Patterson. Primary cultures of dissociated sympathetic neurons. II. Initial studies on catecholamine metabolism. J. Cell Biol. 59: 346–360, 1973.
204.	Mains, R. E., and P. H. Patterson. Primary cultures of dissociated sympathetic neurons. III. Changes in metabolism with age in culture. J. Cell Biol. 59: 361–366, 1973.
205.	Mallart, A., and A. R. Martin. The relation between quantal content and facilitation at the neuromuscular junction of the frog. J. Physiol. London 196: 593–604, 1968.
206.	Marchok, A., and H. Herrmann. Studies of muscle development. I. Changes in cell proliferation. Develop. Biol. 15: 129–155, 1967.
207.	McDonald, T. F., and R. L. DeHaan. Ion levels and membrane potential in chick heart tissue and cultured cells. J. Gen. Physiol. 61: 89–109, 1973.
208.	McDonald, T. F., H. G. Sachs, and R. L. DeHaan. Tetrodotoxin desensitization in aggregates of embryonic chick heart cells. J. Gen. Physiol. 62: 286–302, 1973.
209.	McGeer, P. L., E. G. McGeer, V. K. Singh, and W. H. Chase. Choline acetyltransferase localization in the central nervous system by immunohistochemistry. Brain Res. 81: 373–379, 1974.
210.	McLaughlin, B. J., J. G. Wood, K. Saito, R. Barger, J. E. Vaughn, E. Roberts, and J. Y. Wu. The fine structural localization of glutamate decarboxylase in synaptic terminals of rodent cerebellum. Brain Res. 76: 377–391, 1974.
211.	McMahan, U. J., and S. W. Kuffler. Visual identification of synaptic boutons on living ganglion cells and of varicosities in postganglionic axons in the heart of the frog. Proc. Roy. Soc. London. Ser. B 177: 485–508, 1971.
212.	McMorris, F. A., P. G. Nelson, and F. H. Ruddle (editors)., Contributions of clonal systems to neurobiology. Neurosci. Res. Program Bull. 11: 412–536, 1973.
213.	McMorris, F. A., and F. H. Ruddle. Expression of neuronal phenotypes in neuroblastoma cell hybrids. Develop. Biol. 39: 226–246, 1974.
214.	Mendell, L. M., and E. Henneman. Terminals of single Ia fibers: location, density, and distribution within a pool of 300 homonymous motoneurons. J. Neurophysiol. 34: 171–187, 1971.
215.	Miledi, R. Properties of regenerating neuromuscular synapses in the frog. J. Physiol. London 154: 190–205, 1960.
216.	Miller, R., and R. Phillips. Separation of cells by velocity sedimentation. J. Cellular Physiol. 73: 191–202, 1969.
217.	Minchin, M. C. W., and L. L. Iversen. Release of [3H]gamma‐aminobutyric acid from glial cells in rat dorsal root ganglia. J. Neurochem. 23: 533–540, 1974.
218.	Minna, J. D., Genetic analysis of the mammalian nervous system using somatic cell culture techniques. In: Tissue Culture of the Nervous System, edited by S. Sato. New York: Plenum, 1973, p. 161–185.
219.	Minna, J. D., and A. G. Gilman. Expression of genes for metabolism of cyclic adenosine 3':5'‐monophosphate in somatic cells. II. Effects of prostaglandin E and theophylline on parental and hybrid cells. J. Biol. Chem. 248: 6618–6625, 1973.
220.	Minna, J., D. Glazer, and M. Nirenberg. Genetic dissection of neural properties using somatic cell hybrids. Nature New Biol. 235: 225–231, 1972.
221.	Minna, J., P. G. Nelson, J. Peacock, D. Glazer, and M. Nirenberg. Genes for neuronal properties expressed in neuroblastoma x L‐cell hybrids. Proc. Natl. Acad. Sci. US 68: 234–239, 1971.
222.	Minna, J. D., J. Yavelow, and H. G. Coon. Expression of phenotypes in hybrid somatic cells derived from the nervous system. Genetics 79, Suppl. 2: 373–383, 1975.
223.	Model, P. G., M. B. Bornstein, S. Crain, and G. P. Pappas. An EM study of the development of synapses in cultured fetal mouse cerebellum continuously exposed to xylocain. J. Cell Biol. 49: 362–376, 1971.
224.	Monard, D., F. Solomon, M. Rentsch, and R. Gysin. Glia‐induced morphological differentiation in neuroblastoma cells. Proc. Natl. Acad. Sci. US 70: 1894–1897, 1973.
225.	Morris, J. E. Histogenesis and dependent glutamine synthetase inducibility in embryonic neural retina. Irreversible inhibition of differentiation by 5‐bromodeoxyuridine. Develop. Biol. 35: 125–142, 1973.
226.	Moscona, A. Cell suspensions from organ rudiments of chick embryos. Exptl. Cell Res. 3: 535–539, 1952.
227.	Moscona, A., O. A. Trowell, and E. N. Willmer. Chemical dissociation. In: Cells and Tissues in Culture, edited by E. N. Willmer. New York: Academic, 1965, vol. I, p. 52–56.
228.	Murray, M. R., Nervous tissue in vitro. In: Cells and Tissues in Culture, edited by E. N. Willmer. New York: Academic, 1965, vol. II, p. 373–455.
229.	Murray, M. R., E. R. Peterson, and R. P. Bunge. Some nutritional aspects of myelin sheath formation in cultures of central and peripheral nervous system. Proc. Intern. Congr. Neuropathol., 4th 2: 267, 1962.
230.	Naitoh, Y., R. Eckert, and K. Friedman. A regenerative calcium response in Paramecium. J. Exptl. Biol. 56: 667–681, 1972.
231.	Nakai, J. Dissociated dorsal root ganglia in tissue culture. Am. J. Anat. 99: 81–130, 1956.
232.	Nakai, J., and Y. Kawasaki. Studies on the mechanism determining the course of nerve fibers in tissue culture. I. The reaction of the growth cone to various obstructions. Z. Zellforsch. Mikroskop. Anat. 51: 108–122, 1959.
233.	Narahashi, T. Chemicals as tools in the study of excitable membranes. Physiol. Rev. 54: 813–889, 1974.
234.	Negrete, J., J. del Castillo, I. Escobar, and G. Yankelevich. Correlation between amplitudes and rise times of the miniature endplate potentials in frog muscle. Intern. J. Neurosci. 4: 1–10, 1972.
235.	Nelson, P. G. Nerve and muscle cells in culture. Physiol. Rev. 55: 1–61, 1975.
236.	Nelson, P. G., C. Christian, and M. Nirenberg. Synapse formation between clonal neuroblastoma x glioma hybrid cells and striated muscle. Proc. Natl. Acad. Sci. US 73: 123–127, 1976.
237.	Nelson, P. G., and J. H. Peacock. Electrical activity in dissociated cell cultures from fetal mouse cerebellum. Brain Res. 61: 163–174, 1973.
238.	Nelson, P. G., J. Peacock, and T. Amano. Responses of neuroblastoma cells to iontophoretically applied acetylcholine. J. Cellular Physiol. 77: 353–362, 1971.
239.	Nelson, P. G., J. Peacock, T. Amano, and J. Minna. Electrogenesis in mouse neuroblastoma cells in vitro. J. Cellular Physiol. 77: 337–352, 1971.
240.	Nelson, P. G., B. W. Ruffner, and M. Nirenberg. Neuronal tumor cells with excitable membranes grown in vitro. Proc. Natl. Acad. Sci. US 64: 1004–1010, 1969.
241.	Nicholls, J. G., and D. A. Baylor. Specific modalities and receptive fields of sensory neurons in CNS of the leech. J. Neurophysiol. 31: 740–756, 1968.
242.	Nicholls, J. G., and S. W. Kuffler. Extracellular space as a pathway for exchange between blood and neurons in central nervous system of the leech: ionic composition of glial cells and neurons. J. Neurophysiol. 27: 645–671, 1964.
243.	Nurse, C. A., and P. H. O'Lague. Formation of cholinergic synapses between dissociated sympathetic neurons and skeletal myotubes of rat in cell cultures. Proc. Natl. Acad. Sci. US 72: 1955–1959, 1975.
244.	Obata, K. Transmitter sensitivities of some nerve and muscle cells in culture. Brain Res. 73: 71–88, 1974.
245.	Oger, J., B. G. W. Arnason, N. Pantazis, J. Lehrich, and M. Young. Synthesis of nerve growth factor by L and 373 cells in culture. Proc. Natl. Acad. Sci. US 71: 1554–1558, 1974.
246.	Okun, L. M. Isolated dorsal root ganglion neurons in culture: cytological maturation and extension of electrically active processes. J. Neurobiol. 3: 111–151, 1972.
247.	Okun, L., F. Knight, F. K. Ontkean, and C. A. Thomas. Removal of non‐neuronal cells from suspensions of dissociated embryonic dorsal root ganglia. Exptl. Cell Res. 73: 226–229, 1972.
248.	O'Lague, P. H., K. Obata, P. Claude, E. J. Furshpan, and D. D. Potter. Evidence for cholinergic synapses between dissociated rat sympathetic neurons in cell culture. Proc. Natl. Acad. Sci. US 71: 3602–3606, 1974.
249.	Olmstead, J. B., and G. G. Borissy. Microtubules. Ann. Rev. Biochem. 42: 507–540, 1973.
250.	Olmsted, J. B., and G. G. Borisy. Ionic and nucleotide requirements for microtubule polymerization in vitro. Biochemistry 14: 2996–3005, 1975.
251.	Olson, M. I., and R. P. Bunge. Anatomical observations on the specificity of synapse formation in tissue culture. Brain Res. 59: 19–33, 1973.
252.	O'Neill, M. C., and F. E. Stockdale. A kinetic analysis of myogenesis in vitro. J. Cell Biol. 52: 52–65, 1972.
253.	Oppenheimer, S. B., M. Edidin, C. W. Orr, and S. Roseman. An L‐glutamine requirement for intercellular adhesion. Proc. Natl. Acad. Sci. US 63: 1395, 1969.
254.	Pappas, G. D., E. R. Peterson, E. B. Marvrovsky, and S. M. Crain. Electron microscopy of the in vitro development of mammalian motor end plates. Ann. NY Acad. Sci. 183: 33–45, 1971.
255.	Patrick, J., S. F. Heinemann, J. Lindstrom, D. Schubert, and J. H. Steinbach. Appearance of acetylcholine receptors during differentiation of a myogenic cell line. Proc. Natl. Acad. Sci. US 69: 2762–2766, 1972.
256.	Patterson, B., and J. Prives. Appearance of acetylcholine receptor in differentiating cultures of embryonic chick breast muscle. J. Cell Biol. 59: 241–245, 1973.
257.	Patterson, B., and R. C. Strohman. Myosin synthesis in cultures of differentiating chicken embryo skeletal muscle. Develop. Biol. 29: 113–138, 1972.
258.	Patterson, P. H., and L. L. Y. Chun. The influence of nonneuronal cells on catecholamine and acetylcholine synthesis and accumulation in cultures of dissociated sympathetic neurons. Proc. Natl. Acad. Sci. US 71: 3607–3610, 1974.
259.	Patterson, P., L. Reichardt, and L. Chun. Biochemical studies on the development of primary sympathetic neurons in cell culture. Cold Spring Harbor Symp. Quant. Biol. 40: 389–398, 1975.
260.	Patton, W. D. M., and E. J. Zamis. The pharmacological actions of polymethylene bistrimethylammonium salts. Brit. J. Pharmacol. 4: 381–400, 1949.
261.	Paul, J. Cell and Tissue Culture (4th ed.). Edinburgh: Livingstone, 1970.
262.	Peacock, J. H., F. A. McMorris, and P. G. Nelson. Electrical excitability and chemosensitivity of mouse neuroblastoma x mouse or human fibroblast hybrids. Exptl. Cell Res. 79: 199–212, 1973.
263.	Peacock, J., J. Minna, P. G. Nelson, and M. Nirenberg. Use of aminopterin in selecting electrically active neuroblastoma cells. Exptl. Cell Res. 73: 367–377, 1972.
264.	Peacock, J., and P. G. Nelson. Chemosensitivity of mouse neuroblastoma cells in vitro. J. Neurobiol. 4: 363–374, 1973.
265.	Peacock, H. H., and P. G. Nelson. Synaptogenesis in cell cultures of neurons and myotubes from chickens with muscular dystrophy. J. Neurol. Neurosurg. Psychiat. 36: 389–398, 1973.
266.	Peacock, J. H., P. G. Nelson, and M. W. Goldstone. Electrophysiologic study of cultured neurons dissociated from spinal cords and dorsal root ganglia of fetal mice. Develop. Biol. 30: 137–152, 1973.
267.	Pfeiffer, S. E., Clonal lines of glial cells. In: Tissue Culture of the Nervous System, edited by G. Sato. New York: Plenum 1973, p. 203–230.
268.	Pfeiffer, S. E., H. R. Herschman, J. Lightbody, and G. Sato. Synthesis by a clonal line of rat glial cells of the protein unique to the nervous system. J. Cellular Physiol. 75: 329–340, 1970.
269.	Pfeiffer, S. E., and W. Wechsler. A biochemically differentiated line of Schwann cells. Proc. Natl. Acad. Sci. US 69: 2885–2889, 1972.
270.	Pollak, O. J. Tissue Cultures. New York: Karger, 1969.
271.	Pollard, T. D., and R. R. Weihing. Actin and myosin and cell movement. Chem. Rubber Company Critical Rev. Biochem. 2: 1–66, 1974.
272.	Pomerat, C. M., W. Hendelman, C. Raiborn, Jr., and J. Massey. Dynamic activities of nervous tissue in vitro. In: The Neuron, edited by H. Hydén. New York: Elsevier, 1967, p. 119–178.
273.	Poste, G. Tissue dissociation with proteolytic enzymes. Exptl. Cell Res. 65: 359–367, 1971.
274.	Powell, J. A., and D. M. Fambrough. Electrical properties of normal and dysgenic mouse skeletal muscle in culture. J. Cellular Physiol. 82: 21–38, 1973.
275.	Prasad, K. N. X‐ray‐induced morphological differentiation of mouse neuroblastoma cells in vitro. Nature 234: 471–473, 1971.
276.	Prasad, K. N., and A. W. Hsie. Morphological differentiation of mouse neuroblastoma cells induced in vitro by dibutyryl adenosine 3',5'‐cyclic monophosphate. Nature New Biol. 233: 141–142, 1971.
277.	Prasad, K. N., and B. Mandal. Choline acetyltransferase level in cyclic AMP and X‐ray‐induced morphologically differentiated neuroblastoma cells in culture. Cytobios 8: 75–80, 1973.
278.	Prasad, K. N., and A. Vernadakis. Morphological and biochemical study in X‐ray‐ and dibutyryl cyclic AMP‐induced differentiated neuroblastoma cells. Exptl. Cell Res. 70: 27–32, 1972.
279.	Prives, J. M., and B. M. Patterson. Differentiation of cell membranes in cultures of embryonic chick breast muscle. Proc. Natl. Acad. Sci. US 71: 3208–3211, 1974.
280.	Provine, R. R. Ontogeny of bioelectrical activity in the spinal cord of the chick embryo and its behavioral implications. Brain Res. 41: 365–378, 1972.
281.	Provine, R. R., Neurophysiological aspects of behavior development in the chick embryo. In: Studies on the Development of Behavior and the Nervous System. Behavioral Embryology, edited by G. Gilbert. New York: Academic, 1973, vol. 1.
282.	Rahaminoff, R., and Y. Yaari. Delayed release of transmitter of the frog neuromuscular junctions. J. Physiol. London 228: 241–257, 1973.
283.	Rakic, P., Intrinsic and extrinsic factors influencing the shape of neurons and their assembly into neuronal circuits. In: Frontiers in Neurology and Neuroscience Research, edited by P. Seeman and G. Brown. Toronto: Univ. of Toronto Press, 1974, p. 112–132.
284.	Rall, W. Distinguishing theoretical synaptic potentials computed for different soma‐dendritic distributions of synaptic input. J. Neurophysiol. 30: 1138–1168, 1967.
285.	Rall, W., R. E. Burke, T. G. Smith, P. G. Nelson, and K. Frank. Dendritic location of synapses and possible mechanisms for the monosynaptic EPSP in motoneurons. J. Neurophysiol. 30: 1169–1193, 1967.
286.	Ramon‐Molinar, E., The morphology of dendrites. In: The Structure and Function of Nervous Tissue, edited by G. H. Bourne. New York: Academic, 1968, vol. I, p. 205–267.
287.	Ramon y Cajal, S. Studies on Vertebrate Neurogenesis, translated by L. Guth. Springfield, Ill.: Thomas, 1960.
288.	Ransom, B. R., and P. G. Nelson. Neuropharmacological responses from nerve cells in tissue culture. In: Handbook of Psychopharmacology, edited by L. L. Iverson, S. D. Iverson, and S. H. Snyder. New York: Plenum, 1975, vol. 2, p. 101–127.
289.	Rappaport, C. Trypsinization of monkey kidney tissue. An automatic method for the preparation of cell suspensions. Bull. World Health Organ. 14: 147, 1956.
290.	Rappaport, C, J. P. Poole, and H. P. Rappaport. Studies on properties of surfaces required for growth of mammalian cells in synthetic media. Exptl. Cell Res. 20: 465–510, 1960.
291.	Redfern, P. A. Neuromuscular transmission in newborn rats. J. Physiol. London 209: 701–709, 1970.
292.	Redfern, P. A., and S. Thesleff. Action potential generation in denervated rat skeletal muscle. Part II. The action of tetrodotoxin. Acta Physiol. Scand. 82: 70–78, 1971.
293.	Reese, T. S., and C. Shepherd. Dendro‐dendritic synapses in the central nervous system. In: Structure and Function of Synapses, edited by G. G. Pappas and D. P. Purpura. New York: Raven, 1972, p. 121–136.
294.	Revoltella, R., L. Bertolini, and M. Pediconi. Unmasking of nerve growth factor membrane‐specific binding sites in synchronized murine C1300 neuroblastoma cells. Exptl. Cell Res. 85: 89–94, 1974.
295.	Rinaldini, L. M. The isolation of living cells from animal tissues. Intern. Rev. Cytol. 7: 587–674, 1958.
296.	Ritchie, A. K., and D. M. Fambrough. Electrophysiological properties of the membrane and acetylcholine receptor in developing rat and chick myotubes. J. Gen. Physiol. 66: 327–355, 1975.
297.	Robbins, N., and T. Yonezawa. Developing neuromuscular junctions: first signs of chemical transmission during formation in tissue culture. Science 172: 395–398, 1971.
298.	Robbins, N., and T. Yonezawa. Physiological studies during formation and development of rat neuromuscular junctions in tissue culture. J. Gen. Physiol. 58: 467–481, 1971.
299.	Roisen, F. J., and R. A. Murphy. Neurite development in vitro. II. The role of microfilaments and microtubules in dibutyryl adenosine 3',5'‐cyclic monophosphate and nerve growth factor stimulated maturation. J. Neurobiol. 4: 397–412, 1973.
300.	Rojas, E., and C. Armstrong. Sodium conductance activation without inactivation in pronase perfused axons. Nature New Biol. 229: 117–178, 1971.
301.	Roth, S., and J. Weston. The measurement of intercellular adhesion. Proc. Natl. Acad. Sci. US 58: 974–980, 1967.
302.	Sachs, F., and H. LeCar. Acetylcholine noise in tissue culture muscle cells. Nature New Biol. 246: 214–216, 1973.
303.	Sachs, H. G., T. F. MacDonald, and R. L. DeHaan. Tetrodotoxin sensitivity of cultured embryonic heart cells depends on cell interactions. J. Cell Biol. 56: 255–258, 1973.
304.	Sato, G. (editor)., Tissue Culture of the Nervous System. Current Topics in Neurobiology. New York: Plenum, 1973, vol. 1.
305.	Schon, F., and J. S. Kelly. Autoradiographic localization of [3H]GABA and [3H]glutamate over satellite glial cells. Brain Res. 66: 275–288, 1974.
306.	Schrier, B. K. Surface culture of fetal mammalian brain cells: effect of subculture on morphology and choline acetyl‐transferase activity. J. Neurobiol. 4: 117–124, 1973.
307.	Schrier, B. K., and E. J. Thompson. On the role of the glial cells in the mammalian nervous system. J. Biol. Chem. 249: 1769–1780, 1974.
308.	Schrier, B. K., S. H. Wilson, and M. Nirenberg. Cultured cell systems and methods for neurobiology. In: Methods in Enzymology, edited by S. Fleischer and L. Packer. New York: Academic, 1974, vol. 32, p. 765–788.
309.	Schubert, D., A. J. Harris, C. E. Deving, and S. Heinnemann. Characterization of a unique muscle cell line. J. Cell Biol. 61: 398–413, 1974.
310.	Schubert, D., A. J. Harris, S. Heinemann, Y. Kidokoro, J. Patrick, and J. H. Steinbach. Differentiation and interaction of clonal cell lines of nerve and muscle. In: Tissue Culture of the Nervous System, edited by G. Sato. New York: Plenum, 1973, p. 55–86.
311.	Schubert, D., S. Heineman, W. Carlisle, H. Tarikas, B. Kimes, J. Patrick, J. H. Steinbach, W. Culp, and B. L. Brandt. Clonal cell lines from the rat central nervous system. Nature 249: 224–227, 1974.
312.	Schubert, D., S. Humphreys, C. Baroni, and M. Cohen. In vitro differentiation of a mouse neuroblastoma. Proc. Natl. Acad. Sci. US 64: 316–323, 1969.
313.	Schubert, D., S. Humphreys, F. Jacob, and F. DeVitry. Induced differentiation of a neuroblastoma. Develop. Biol. 25: 514–546, 1971.
314.	Schubert, D., and F. Jacob. 5‐Bromodeoxyuridine‐induced differentiation of a neuroblastoma. Proc. Natl. Acad. Sci. US 67: 247–254, 1970.
315.	Scott, B. S. Effect of potassium on neuron survival in cultures of dissociated human nervous tissue. Exptl. Neurol. 39: 297–308, 1971.
316.	Scott, B. S., and K. C. Fisher. Potassium concentration and number of neurons in cultures of dissociated ganglia. Exptl. Neurol. 27: 16–22, 1970.
317.	Scott, B. S., and K. C. Fisher. Effect of choline, high potassium, and low sodium on the number of neurons in cultures of dissociated chick ganglia. Exptl. Neurol. 31: 183–188, 1971.
318.	Seecoff, R. L., N. Alleaume, R. L. Teplitz, and I. Gerson. Differentiation of neurons and myocytes in cell cultures made from Drosophila gastrulae. Exptl. Cell Res. 69: 161–173, 1971.
319.	Seeds, N. Biochemical differentiation in reaggregating brain cell culture. Proc. Natl. Acad. Sci. US 68: 1858–1861, 1971.
320.	Seeds, N. W., Differentiation of aggregating brain cell cultures. In: Tissue Culture of the Nervous System, edited by G. Sato. New York: Plenum, 1973, p. 35–53.
321.	Seeds, N. W., A. Gilman, T. Amano, and M. Nirenberg. Regulation of axon formation by clonal lines of a neural tumor. Proc. Natl. Acad. Sci. US 66: 160–167, 1970.
322.	Seravin, L. N. Mechanisms and coordination in cellular locomotion. Advan. Comp. Physiol. Biochem. 4: 37–111, 1971.
323.	Shainberg, A., G. Yagil, and D. Yaffe. Control of myogenesis by Ca++ concentration in nutritional medium. Exptl. Cell Res. 58: 163–166, 1969.
324.	Shainberg, A., G. Yagil, and D. Yaffe. Alterations of enzyme activities during muscle differentiation in vitro. Develop. Biol. 25: 1–19, 1971.
325.	Sheridan, J., Electrical coupling of cells and cell communication. In: Intercellular Communication, edited by R. Cox. New York: Wiley, 1974, p. 30–42.
326.	Shimada, Y. Suppression of myogenesis by heterotypic and heterospecific cells in monolayer cultures. Exptl. Cell Res. 51: 564–578, 1968.
327.	Shimada, Y., and D. A. Fischman. Morphological and physiological evidence for the development of functional neuromuscular junctions in vitro. Develop. Biol. 31: 200–225, 1973.
328.	Shimada, Y., D. A. Fischman, and A. A. Moscona. Formation of neuromuscular junctions in embryonic cell cultures. Proc. Natl. Acad. Sci. US 62: 715–721, 1969.
329.	Shimada, Y., D. A. Fischman, and A. A. Moscona. The development of nerve‐muscle junctions in monolayer cultures of embryonic spinal cord and skeletal muscle cells. J. Cell Biol. 43: 382–387, 1969.
330.	Sidman, R., Autoradiographic methods and principles for study of the nervous system with thymidine‐H3. In: Contemporary Research Methods in Neuroanatomy, edited by W. S. H. Nata and S. O. E. Ebbesson. New York: Springer Verlag, 1970, p. 252–274.
331.	Simantov, R., and L. Sachs. Enzyme regulation in neuroblastoma cells. Selection of clones with low acetylcholinesterase activity and the independent control of acetylcholinesterase and choline‐O‐acetyltransferase. European J. Biochem. 30: 123–129, 1972.
332.	Simantov, R., and L. Sachs. Regulation of acetylcholine receptors in relation to acetylcholinesterase in neuroblastoma cells. Proc. Natl. Acad. Sci. US 70: 2902–2905, 1973.
333.	Skoff, R. P., and V. Hamburger. Fine structure of dendritic and axonal growth cones in embryonic chick spinal cord. J. Comp. Neurol. 153: 107–148, 1974.
334.	Smilowitz, H., R. Siegel, and G. D. Fischbach. Freshly dissociated myoblasts contain ACh receptors. J. Cell Biol. 63: 320a, 1974.
335.	Sobkowicz, H. M., H. A. Hartmann, R. Monzain, and P. Desnoyers. Growth, differentiation and ribonucleic acid content of the fetal rat spinal ganglion cells in culture. J. Comp. Neurol. 148: 249–284, 1973.
336.	Spector, I., Y. Kimhi, and P. G. Nelson. Tetrodotoxin and cobalt blockade of neuroblastoma action potentials. Nature New Biol. 246: 124–126, 1973.
337.	Speidel, C. Studies of living nerves. II. Activities of ameboid growth cones, sheath cells, and myelin segments as revealed by prolonged observation of individual nerve fibers in frog tadpoles. Am. J. Anat. 52: 1–79, 1933.
338.	Sperelakis, N., and K. Shigenabu. Changes in membrane properties of chick embryonic hearts during development. J. Gen. Physiol. 60: 430–453, 1972.
339.	Spooner, B. S., K. M. Yamada, and N. K. Wessells. Microfilaments and cell locomotion. J. Cell Biol. 49: 595–613, 1971.
340.	Stambrook, P. J., H. G. Sachs, and J. D. Ebert. The effect of K+ on cell membrane potential and the passage of cells through the cell cycle: a block in G1. In: Carnegie Institution Year Book, 1972, p. 64–69.
341.	Steinbach, J. H. Acetylcholine responses on clonal myogenic cells in vitro. J. Physiol. London 247: 393–405, 1975.
342.	Steinbach, J. H., A. J. Harris, J. Patrick, D. Schubert, and S. Heinemann. Nerve‐muscle interaction in vitro. The role of acetylcholine. J. Gen. Physiol. 62: 255–270, 1973.
343.	Stewart, H. L., K. C. Sneill, L. J. Dunham, and S. M. Schlyen. Transplantable and transmissible tumors of animals. In: Atlas of Tumor Pathology. Washington, D.C.: Armed Forces Inst. Pathol. Natl. Acad. Sci. 1959, sect. XII, fasc. 40.
344.	Stillwell, E. F., C. M. Cone, and C. M. Cone, Jr. Stimulation of DNA synthesis in CNS neurones by sustained depolarisation. Nature New Biol. 246: 110–111, 1973.
345.	Stockel, K., U. Paravicini, and H. Thoenen. Specificity of the retrograde axonal transport of nerve growth factor. Brain Res. 76: 413–421, 1974.
346.	Stockel, K., F. Solomon, U. Paravicini, and H. Thoenen. Dissociation between effects of nerve growth factor on tyrosine hydroxylase and tubulin synthesis in sympathetic ganglia. Nature 250: 150–151, 1974.
347.	Stykowski, A. J., Z. Vogel, and M. W. Nirenberg. Development of acetylcholine receptor clusters on cultured muscle cells. Proc. Natl. Acad. Sci. US 70: 270–274, 1973.
348.	Takahashi, K., S. Miyazaki, and Y. Kidokoro. Development of excitability in embryonic muscle cell membranes in certain tunicates. Science 171: 415–417, 1971.
349.	Tello, J. F. Genesis de las terminaniones nervidas moterices y sensitivas I. En la sistema locomotor de las vergebrados superiores. Histogenesis muscular. Trabajos Inst. Cajal Invest. Biol. Madrid 15: 101–199, 1917.
350.	Tennyson, V. M. The fine structure of the axon and growth cone of the dorsal root neuroblast of the rabbit embryo. J. Cell Biol. 44: 62–79, 1970.
351.	Tobias, J. M., and P. G. Nelson. Structure and function in nerve. In: A Symposium on Molecular Biology, edited by R. Zirkle. Chicago: Univ. of Chicago Press, 1959, p. 248–265.
352.	Trinkaus, J. P. Cells into Organs: the Forces That Shape the Embryo. Englewood Cliffs, N. J.: Prentice‐Hall, 1969.
353.	Van Essen, D., and J. K. S. Jansen. Reinnervation of the rat diaphragm during perfusion with α‐bungarotoxin. Acta Physiol. Scand. 91: 571–573, 1974.
354.	Varon, S. S., and C. W. Raiborn, Jr. Dissociation, fractionation and culture of embryonic brain cells. Brain Res. 12: 180–199, 1969.
355.	Varon, S., and C. W. Raiborn, Jr. Excitability and conduction in neurons of dissociated ganglionic cell cultures. Brain Res. 30: 83–89, 1971.
356.	Varon, S., and C. Raiborn. Dissociation, fractionation and culture of chick embryo sympathetic ganglion cells. J. Neurobiol. 1: 211–221, 1972.
357.	Vaughn, J. E., C. K. Henrikson, and J. A. Greishaber. A quantitative study of synapses on motor neuron dendritic growth comes in developing mouse spinal cord. J. Cell Biol. 60: 664–672, 1974.
358.	Vernadakis, A., and D. M. Woodbury. Electrolyte and nitrogen changes in skeletal muscles of developing rats. Am. J. Physiol. 206: 1365–1368, 1964.
359.	Vogel, Z., A. J. Sytkowski, and M. W. Nirenberg. Acetylcholine receptors of muscle grown in vitro. Proc. Natl. Acad. Sci. US 69: 3180–3184, 1972.
360.	Wald, F. Ionic differences between somatic and axonal action potentials in snail giant neurons. J. Physiol. London 220: 267–281, 1972.
361.	Wallach, D. F., and P. S. Lin. A critical evaluation of plasma membrane fractionation. Biochim. Biophys. Acta 300: 211–254, 1973.
362.	Warters, R. L., and K. G. Hofer. The in vivo reproduction potential of density separated cells. Exptl. Cell Res. 87; 143–151, 1974.
363.	Waymouth, C., The cultivation of cells in chemically defined medium and the malignant transformation of cells in vitro. In: Tissue Culture, edited by C. V. Ramakrishnan. The Hague: W. Junk, 1965, p. 168–179.
364.	Waymouth, C. To disaggregate or not to disaggregate. Inquiry and cell disaggregation, transient or permanent. In Vitro 10: 97–111, 1974.
365.	Wechsler, W., S. E. Pfeiffer, J. A. Swenberg, and A. Koestner. S‐100 protein in methyl‐ and ethylnitrosourea induced tumors of the rat nervous system. Acta Neuropathol. Berlin 24: 287–303, 1973.
366.	Weiss, P. In vitro experiments on the factors determining the course of the outgrowing nerve fiber. J. Exptl. Zool. 68: 393–448, 1934.
367.	Weiss, P. Experiments on cell and axon orientation in vitro: the role of colloidal exudates in tissue organization. J. Exptl. Zool. 100: 353–386, 1945.
368.	Wessels, N. K., B. S. Spooner, J. F. Ash, M. O. Bradley, M. O. Luduena, E. L. Taylor, J. T. Wrenn, and K. M. Yamada. Microfilaments in cellular and developmental processes. Science 171: 135–143, 1971.
369.	Wessels, N., B. Spooner, and M. Ludena. Surface movements, microfilaments and cell locomotion. In: Locomotion of Tissue Cells. New York: Elsevier, 1973, 53–82. (Ciba Foundation Symp. 14.)
370.	Wickus, G., E. Gruenstein, P. Robbins, and A. Rich. Decrease in membrane‐associated actin of fibroblasts after transformation by Rous sarcoma virus. Proc. Natl. Acad. Sci. US 72: 747–749, 1975.
371.	Willmer, E. N. Tissue Culture; The Growth and Differentiation of Normal Tissues in Artificial Media (3rd ed.). New York: Wiley, 1958.
372.	Willmer, E. N. (editor)., Cells and Tissues in Culture. New York: Academic, 1965, vols. I‐III.
373.	Wilson, B. W., P. S. Nieberg, C. R. Walker, T. A. Linkhart, and D. M. Fry. Production and release of acetylcholine esterase by cultured chick embryo muscle. Develop. Biol. 33: 285–289, 1973.
374.	Wilson, S. H., B. K. Schrier, J. L. Farber, E. J. Thompson, R. N. Rosenberg, A. J. Blume, and M. W. Nirenberg. Markers for gene expression in cultured cells from the nervous system. J. Biol. Chem. 247: 3159–3169, 1972.
375.	Yaffe, D. Retention of differentiation potentialities during prolonged cultivation of myogenic cells. Proc. Natl. Acad. Sci. US 61: 477–483, 1968.
376.	Yaffe, D., Cellular aspects of muscle differentiation in vitro. In: Current Topics in Developmental Biology, edited by A. A. Moscona and A. Monroy. New York: Academic, 1969, vol. 4, p. 37–77.
377.	Yaffe, D. Developmental changes preceding cell fusion during muscle differentiation in vitro. Exptl. Cell Res. 66: 33–48, 1971.
378.	Yamada, K. M., B. S. Spooner, and N. K. Wessels. Axon growth: roles of microfilaments and microtubules. Proc. Natl. Acad. Sci. US 66: 1206–1212, 1970.
379.	Yamada, K. M., B. S. Spooner, and N. K. Wessells. Ultrastructure and function of growth cones and axons of cultured nerve cells. J. Cell Biol. 49: 614–635, 1971.
380.	Yamada, K. M., and N. K. Wessells. Axon elongation. Effect of nerve growth factor on microtubule protein. Exptl. Cell Res. 66: 346–352, 1971.
381.	Yavin, E., and Z. Yavin. Attachment and culture of dissociated cells from rat embryo cerebral hemispheres on polylysine coated surface. J. Cell Biol. 62: 540–546, 1974.

Contact Editor

Submit a note to the editor about this article by filling in the form below.

* Required Field

Article Title:	Cell Culture in Neurobiology
* Name:
* E-mail:
* Note:

How to Cite

Gerald D. Fischbach, Phillip G. Nelson. Cell Culture in Neurobiology. Compr Physiol 2011, Supplement 1: Handbook of Physiology, The Nervous System, Cellular Biology of Neurons: 719-774. First published in print 1977. doi: 10.1002/cphy.cp010120

Collections

Free Featured Articles