Evidence for Four Classes of Microtubules in Individual Cells

J. Cell Sci. 2, 169-192 (1967) 169 Printed in Great Britain

EVIDENCE FOR FOUR CLASSES OF MICROTUBULES IN INDIVIDUAL CELLS

O.BEHNKE Department of Anatomy, Royal Dental College, Universitetsparken 4, Copenhagen 0, Denmark AND A. FORER# Carlsberg Foundation, Biological Institute, Tagensvej 16, Copenhagen N, Denmark

SUMMARY Experiments were performed on crane-fly spermatids (Nephrotoma suturalis Loew), rat- sperm, and rat tracheal cilia to test whether all microtubules respond in the same way to different treatments. Crane-fly spermatids contain cytoplasmic microtubules, accessory tubules, and the 9 + 2 complex of tubules; rat sperm and rat tracheal cilia contain only the 9 + 2 tubules. Crane-fly spermatid tubules responded to the experimental treatments as follows. After colchicine treatment, or storage at o°C, the cytoplasmic microtubules disappeared, while the 9 + 2 tubules were normal. After storage at 50 CC the cytoplasmic microtubules disappeared, and then the 9 + 2 tubules were affected: first the central tubules and B-tubules were affected, and later the A-tubules. After brief pepsin treatment, the 9 doublet tubules disappeared, while the other tubules appeared normal; after prolonged pepsin treatment the accessory, central, and cytoplasmic tubules disappeared. After negative staining at pH 7, the cytoplasmic microtubules were never seen, the central tubules were only sometimes seen, the B-tubules were sometimes fragmented, and the A-tubules were intact. On the basis of these responses, it was concluded that there are 4 classes of tubules in crane-fly spermatids, namely cytoplasmic microtubules; accessory tubules and central tubules (of the 9 + 2 complex); B-tubules (of the 9 + 2 complex); and A-tubules (of the 9 + 2 complex). At least some of the different responses appeared to be due to intrinsic physical and/or chemical differences between the tubules themselves. Pepsin digestion and negative staining of rat sperm tails gave results similar to those with crane-fly spermatids. In addition, the 9 + 2 tubules responded differently to pepsin digestion at different points along their length. This gradient of sensitivity was attributed to synthesis of new tubules occurring at one end of the sperm tail. Pepsin digestion and negative staining of rat tracheal cilia gave results similar to those with crane-fly spermatids and rat sperm tails. All the tubules had a similar substructure, as revealed by negative-staining techniques. It was concluded that microtubules are proteinaceous, at least in part, and that microtubules are different in composition from membranes. It is suggested that the walls of the B-tubules are composed of two materials—(1) the portions adjacent to the A-tubules, and (2) the remaining portion.

INTRODUCTION The term ' microtubule' is currently used to designate cylindrical cellular structures with electron-dense walls and less dense cores, and with an outer diameter ranging from about 180 to 300 A (Slautterback, 1963; Ledbetter & Porter, 1963). Micro- tubules have been found in the cytoplasm of almost all cell types studied (see Slautter- • Present address: Department of Zoology, Downing Street, Cambridge 170 O. Behnke and A. Forer back, 1963; Pitelka, 1963; Byers & Porter, 1964; Silveira & Porter, 1964; Behnke & Forer, 1966a). They are universally associated with spermatid tail9, 9perm tails, cilia, flagella, and mitotic spindles. Microtubules are generally straight and can often be followed for distances of microns, though sometimes they are evenly curved (Fawcett & Witebsky, 1964; Behnke, 19656; Haydon & Taylor, 1965; Hoffman, 1966) or wavy (Vivier & Andre", 1961; Barnicot & Huxley, 1965; Carasso & Favard, 1965; Newcomb & Bonnett, 1965; Vivier, 1965; Behnke & Forer, 19666). They can sometimes be identified by the patterns in which they are arranged, or the organelles with which they are associated. For example, microtubules arranged in a consistent '9 + 2' pattern are regularly found in cilia and flagella, and in axial filaments of spermatids and sperm (Manton & Clarke, 1952; Fawcett & Porter, 1954; Afzelius, 1959; Gibbons & Grimstone, i960; Fawcett, 1961). Other consistent patterns of microtubules are found in other cells (see, for example, Tilney & Porter, 1965; Batisse, 1965; Lumsden, 1965; Grimstone & Gibbons, 1966; Nilsson & Williams, 1966). Some microtubules are not arranged in such a precise way, but can nonetheless be identified because they are located in specific regions of the cell, such as near nuclei (Burgos & Fawcett, 1955), in axostyles (Grimstone & Cleveland, 1965), in spindles (Harris, 1962; Kane, 1962; Roth & Daniels, 1962; Dales, 1963; Ledbetter & Porter, 1963), etc. For lack of other distinguishing characteristics, microtubules which are not arranged in any specific pattern and are not located in any specific region are designated ' cytoplasmic microtubules'. The function of microtubules is not known. From studies of sectioned material it has been suggested that microtubules are involved in cell motility, or are involved in cytoplasmic streaming, or provide skeletal support, or transport water and small ions. There is, however, no conclusive evidence to support these suggestions, nor is it known if all microtubules are equivalent, or have the same function. Some workers (for example, Slautterback, 1963; H. Moor, unpublished observations) have tried to classify microtubules by size, but because of technical and biological variations this idea has not been generally accepted. Some have suggested a classification by fixation properties (Behnke & Zelander, 1966; Sheffield, 1966), to account for the observation that some microtubules are well preserved with osmium tetroxide fixation while others require special fixation, such as glutaraldehyde, or low pH of the osmium fixative together with addition of cations (Harris, 1962; Roth & Daniels, 1962). In general, however, it is thought that all microtubules are the same. Because of their morphological similarities, various authors have suggested that tubules in cilia and flagella may be the same as cytoplasmic microtubules (Ledbetter & Porter, 1963; Bawa, 1964; Byers & Porter, 1964; Porter, Ledbetter & Badenhausen, 1964; Silveira & Porter, 1964; Anderson, Weissmann & Ellis, 1966; Robison, 1966) or the same as spindle microtubules (Ledbetter & Porter, 1963; Pease, 1963; Harris & Bajer, 1965; Krishan & Buck, 1965; Kiefer, Sakai, Solari & Mazia, 1966), and also that spindle microtubules are the same as cytoplasmic microtubules (Ledbetter & Porter, 1963; de-The", 1964; Anderson et al. 1966). We report here experiments designed to show whether or not all microtubules are Classes of microtubules 171 the same. We conclude that they are not, because various experimental treatments produce different reactions with different microtubules. The results show: there are at least 4 classes of microtubules; within at least 3 of the classes the microtubules differ along their length; microtubules have a similar substructure, as revealed by negative staining techniques; and microtubules are proteinaceous, at least in large part, and their walls are different in composition from cell membranes. Further data suggest that different microtubules contain different material in their less-dense cores, and that the walls of some individual tubules have two components.

MATERIALS AND METHODS Crane-fly testes Crane flies (Nephrotoma suturalis Loew) were reared in the laboratory. Last-instar larvae were dissected at a time when their testes contained meiotic cells and young spermatids, or contained spermatids 1-4 days after the second meiotic division (see Behnke & Forer (19666) or Forer (1964, 1965) for details of the animals and spermatocytes). Larvae were dissected at room temperature under Kel-F 10 oil (Minnesota Mining and Mfg Co.), which prevents dehydration of the testes during dissection. Individual testes (and a surrounding film of oil) were either placed directly into fixative, at room temperature, and processed for electron microscopy, or transferred to an experimental solution. In both cases, the testes sank under the surface of the solution as the surrounding film of oil floated on top. Fixation. Except for experiments in which different fixation techniques were studied, all testes were fixed in glutaraldehyde and processed as follows. Testes were fixed by immersion in 2-4 % glutaraldehyde solution ino-iM cacodylate buffer, at pH between pH 6-5 and 7-4. They were post-fixed for 1 h in 1 % osmium tetroxide solution (in o-i M cacodylate or veronal-acetate buffer, at pH 7), and then dehydrated through a series of alcohols and embedded in Epon (Luft, 1961). Thin sections were double stained, with uranyl acetate (Watson, 1958) and lead citrate (Reynolds, 1963), and examined in a Siemens Elmiskop I. Some testes were fixed at room temperature in 1 % osmium tetroxide solution, which was prepared either in o-i M cacodylate buffer at pH 7, with CaCl, added to a finalconcentratio n of Ca1+ of 0-002 M, or in o-i M cacodylate buffer at pH 6-5 without Ca1+ added, or in veronal- acetate buffer, at pH 7^4, without Ca1+. Experimental treatments of testes. Some testes were exposed to different temperatures. These were placed in insect Ringer solution (Ephrussi & Beadle, 1936) which was previously equili- brated at the experimental temperature, 0°, 40°, or 50° C. At various times (10-30 min) an equal volume of 4 % glutaraldehyde solution at room temperature was added, and the fixation continued for 2 h at room temperature. The testes were then processed as described above, for non-treated testes. In some experiments testes were transferred from the experimental Ringer solution to fixative at room temperature. Some testes were treated with colchicine. These were placed in Ringer solution at room temperature, to which colchicine had been added to a final concentration of I^XIO^M. They were removed after various times, fixed with glutaraldehyde and processed as above. Control testes were placed in Ringer solution and kept at room temperature for 60 min (the maximum experimental treatment) prior to fixation. Pepsin digestion of sections from non-treated testes. Enzyme digestions were performed on sections of non-treated testes, in principle according to the method of Bernhard, Granboulan, Barski & Tournier (1961). Sections of glutaraldehyde-osmiurn-fixed Epon-embedded crane-fly testes were placed on grids. The latter were then floated for 10 min on a drop of 10 % H,O, solution (to oxidize the reduced osmium fixative to soluble osmium tetroxide), thoroughly rinsed in distilled water, and, at 37 °C in a moist chamber, floated for various times on a drop of 0-5 % pepsin (Difco, 1:10000) in o-i N HC1. Digestion was carried out for various times from 5-30 min. After digestion, the sections were rinsed with distilled water, and finallystaine d with uranyl acetate and lead citrate. 172 O. Behnke and A. Forer

Control sections were treated with H2O| and floated for 30 min on o-i N HC1. Negative staining. Testes were placed in a drop of distilled water, or insect Ringer solution, and transferred through several such drops to remove the adhering Kel-F 10 oil. The testes. were pierced on a drop of distilled water, insect Ringer solution, or the negative stain solution. After piercing, the testicular contents spread themselves on the surface of the drop. The material was picked up by touching carbon-coated, Formvar-covered grids to the surface. Excess liquid was drained off by touching the edge of the grid to filter paper, and (except for material lysed directly in stain) the adhering material was negatively stained with freshly prepared 1 % potassium phosphotungstate solution, at pH 4, 5 or 7. Material which was lysed in the negative stain was used directly after excess liquid had been drained off. Some material was shadowed with platinum-carbon.

Rat testes and rat tracheal cilia Fixation for sections. Pieces of testes were removed from young adult albino rats, and fixed by immersion for 2 h in 3 % glutaraldehyde solution, at pH 7 in o-1 M cacodylate buffer. They were post-fixed for 1 h in 1 % osmium tetroxide solution in o-i M cacodylate buffer (pH 7), and processed further as described above for crane-fly testes. The lungs and trachea were removed from rats after 3 % glutaraldehyde solution (at pH 7, ino-i M cacodylate buffer) was placed in the trachea. The tissue was then cut into pieces, fixed by immersion for 2 h in 3 % glutaraldehyde solution, and processed further as described for rat testes. Negative staining. Rat sperm were removed from the epididymis and placed on a drop of distilled water. Cilia were obtained from rat trachea by gently scraping the surface of the epithelial lining with a sharp instrument, and the material placed on the surface of a drop of distilled water. The cells or cilia were picked up from the drop of water on to carbon-coated, Formvar- covered grids, and negatively stained with 1 % potassium phosphotungstate at pH 5. Pepsin digestion of sections of normal tissue. Pepsin digestions were performed as described for crane-fly spermatids.

RESULTS Crane-fly spermatids and spermatocytes Normal morphology. Glutaraldehyde-fixed spermatids always contained at least one 9 + 2 complex of tubules (axial filament tubules) in the tail, and often there were several; as many as 9 have been observed in one spermatid. The 9 + 2 tubules did not differ from the usual descriptions (Afzelius, 1959; Gibbons & Grimstone, i960; Fawcett, 1961). 'Spokes' (Afzelius, 1959) radiated from the region of the 2 central tubules ('fibres', of Gibbons & Grimstone, i960) to the 'A' member of each of the 9 doublets (Figs. 3-5). The A-tubules ('subfibres', of Gibbons & Grimstone, i960) had 'arms'; the B-tubules, which were not completely tubular, did not (Figs. 3, 4). The doublets were at a slight angle to the tangent of the circle, so that the A-tubules were always closer to the centre than the B-tubules (Figs. 3, 4), similar to the situation described by Gibbons & Grimstone (i960). At least one extremely long mitochondrion was found in each spermatid, usually located near the 9 + 2 tubules and apparently extending almost the entire length of the sperm tail. Both the mitochondrion and the 9 + 2 tubules were usually associated with a membrane system (endoplasmic reticulum) which also extended almost the full length of the tail. In more mature spermatids the interior of the A-tubules was somewhat electron- Classes of microtubules 173 dense (Fig. 4), and there was an accessory microtubule associated with each of the 9 doublets (Fig. 4), similar to the accessory microtubules described by others (Kaye, 1964; Bawa, 1964; Cameron, 1965). In younger spermatids there were many cytoplasmic microtubules throughout the •cytoplasm, parallel to the 9 + 2 tubules (Fig. 3). The cytoplasmic microtubules were not associated with any particular organelle or region. In more mature spermatids, the •cytoplasmic microtubules were still oriented parallel to the 9 + 2 tubules, but many of them were associated with the mitochondrion (Fig. 4). However, in these spermatids there were always some cytoplasmic microtubules which were not associated with the mitochondrion, or any other organelle or region. It is difficult to estimate the numbers of cytoplasmic microtubules, especially since the numbers seen depend on the stage of differentiation, and the region where the tail is sectioned. To give a rough idea of the numbers seen in sections of normal spermatids: young spermatids (1 day after meiosis) contained of the order of 20—50 cytoplasmic microtubules, generally not associated with any particular region (Fig. 3), while more mature spermatids (3-4 days after meiosis) contained of the order of 10—50, varying numbers of which were localized near the mitochondrion (Fig. 4). In sectioned material, cytoplasmic microtubules often seemed to have a substructure. In cross-sections an electron-dense line often extended between 2 cytoplasmic microtubules, joining them in pairs (Figs. 3, 4), as has also been noted for microtubules in Giardia muris (Friend, 1966). A similar line was often seen extending from some of the accessory microtubules toward the doublets (Fig. 4), similar to lines on accessory tubules described by Bawa (1964) and Cameron (1965). The cytoplasmic microtubules, the accessory tubules, the central microtubules •of the 9 + 2 complex and the A-tubules were all about 250 A in diameter. Osmium-tetroxide-fixed spermatids appeared somewhat different from the above description. All three solutions of osmium tetroxide used gave the same results: the 9 + 2 and accessory tubules were present in spermatids, and these tubules were well preserved; that is, they looked the same as in glutaraldehyde-fixed spermatids. Cytoplasmic microtubules, however, were not consistently seen in osmium-tetroxide- fixed spermatids, and when they were present they were not well preserved; that is to say, their outlines were not well demarcated, and they often appeared elliptical and/or broken. Frequently they were quite ill-defined and distorted, so that if cytoplasmic microtubules had not been seen regularly in glutaraldehyde-fixed material it would have been difficult to be sure of their presence. In osmium-fixed spermatids, the A-tubules and central tubules (of the 9 + 2 complex) were about 250 A in diameter, and the cytoplasmic tubules, when present, were about the same. Crane-fly spermatocytes during division have been described elsewhere (Behnke & Forer, 19666). These dividing cells contain both spindle microtubules and cytoplasmic microtubules. Morphology of experimentally treated cells. Spermatids stored at o °C for 15 min contained normal 9 + 2 and accessory tubules, but usually no cytoplasmic microtubules. Occasionally, a few of the latter could be seen (Figs. 5, 6). Thus, cytoplasmic tubules are more sensitive to cold treatment of cells than the 9 + 2 and accessory 174 O- Behnke and A. Forer tubules. Dividing spermatocyte9 stored at o °C for 15 min contained neither spindle nor cytoplasmic microtubules. Crane-fly spermatids stored at 50 °C for 10 min contained the 9 + 2 tubules, but no cytoplasmic microtubules. Longer times (15-25 min) at 50 °C resulted in changes in the morphology of the 9 + 2 tubules. The central and B-tubules were affected first, and the A-tubules last. The central tubules disappeared; affected cells contained either one (Fig. 8) or none. Sometimes there was an indication of a fragment of tubule remaining (an arc or line in cross-section). Usually, however, the tubule(s) had disappeared completely. B-tubules at first were separated from the A-tubules at the central side of the A-B junction; such doublets often appeared in cross-section like the numeral 6 (Figs. 8, 9). (The arms cannot be identified after treatment at 50 °C, but the disarranged tubule could be distinguished as the B-tubule, because the unaffected member was tubular, was associated with the spokes, and was found closest to the centre of the complex (Figs. 8, 9).) Later, the B-tubules disintegrated into fragments (Fig. 8). Finally, all the B-tubules disappeared, and the 9 + 2 complex was reduced to a circle of A-tubules (Fig. 10). Central and B-tubules were affected at about the same time, but were apparently affected independently, for sometimes some B-tubules were affected when the central tubules were normal (Fig. 9), and conversely. The A-tubules were affected only after the central and B-tubules had disappeared. The ring of A-tubules became disordered (Fig. 11), and eventually the A-tubules disappeared, as shown by the fact that many spermatids contained no tubules at all. This sequence of events is reconstructed from the various abnormalities observed in the 9 + 2 tubules after treatment at 50 °C. These abnormalities are illustrated diagrammatically in Fig. 1, together with our interpretation of the sequence of events. We should emphasize that this description of the various disarrangements is not quantitative, for in individual sections we often observed spermatids showing more than one stage of the described abnormalities. The results of treatment at 50 °C can thus be summarized as: cytoplasmic microtubules > (central tubules) = (B-tubules) > (A-tubules), where the notation X > Y means that X is more sensitive to the treatment than Y. Spermatids kept at 40 °C for 25 min were not different from the controls. Storage of spermatocytes at 50 °C for 15 min had drastic effects on the general morphology of the cells. Chromosomes were recognizable, but neither spindle microtubules nor cytoplasmic microtubules could be identified. Colchicine, too, affected the cytoplasmic microtubules in spermatids. After 20 min in colchicine all crane-fly spermatids contained normal 9 + 2 tubules, and some spermatids contained a few cytoplasmic microtubules. After 40 and 60 min in colchicine, the 9 + 2 tubules were normal, but all cytoplasmic microtubules had disappeared (Fig. 7). Only a few observations were made on colchicine-treated spermatocytes; though preliminary, they are worth mentioning. Normally, dividing crane-fly spermatocytes contain spindle microtubules which pass through chromosomes, spindle microtubules not associated with chromosomes, microtubules in the mitochondrial sheath surrounding the spindle, and microtubules in the cytoplasm outside the mitochondrial sheath Classes of microtubules 175 (Behnke & Forer, 19666). After 40-60 min in colchicine the spindle birefringence was gone and the chromosomes were scattered throughout the cell, but at least some of the microtubules which pass through the chromosomes were still present, enclosed within the chromosomes, and at least some of the microtubules associated with the mitochondria were still present; few other microtubules were seen. Since in normal

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12 Cell Sci. 2 178 O. Behnke and A. Forer about 30 A in diameter, spaced (centre to centre) about 45 A apart (Fig. 25). This beaded appearance seemed to be related to the pH of the stain; it was observed more often when the negative stain was at pH 7 than at pH 5 or below. Beading was more common in 9 + 2 tubules than in cytoplasmic tubules, but this may be because cytoplasmic tubules were not seen at pH 7 (see below). The different kinds of tubules differed in stability during the staining. When the stain was at pH 7 only the 9 + 2 tubules were seen, and, of these, the central tubules were rarely observed. When the central tubules were present, they often did not remain as intact tubules, but rather the individual filaments were separated and frayed out. The B-tubules were in a few cases partially fragmented and frayed into filaments, while the A-tubules were generally intact. At pH 7, then, the A-tubules were the most stable, the B-tubules next, and the central tubules least. The cytoplasmic tubules seemed to be completely unstable, since none was seen. When the negative stain was at pH 5 cytoplasmic microtubules were seen, as well as the 9 + 2 tubules (Figs. 26, 27). Therefore we conclude that cytoplasmic microtubules are more stable at pH 5 than 7. Summary of experiments on crane-fly spermatid microtubules. The experiments on crane-fly spermatid microtubules are summarized in Table 1. On the basis of the responses to the various treatments it is possible to distinguish between four classes of tubules in this one cell, namely, (1) cytoplasmic microtubules, (2) central tubules and accessory tubules, (3) B-tubules, and (4) A-tubules. On the basis of limited informa- tion, spindle microtubules would tentatively be in class (1).

Rat testes Morphology of normal cells. The morphology of normal rat sperm tails has been described by others (Telkka, Fawcett & Christensen, 1961), and no detailed description will be given here. It is relevant to point out, however, that one can determine the position of a cross-section along the length of the sperm by determining whether the axial complex is associated with 'coarse bodies' (mid-piece), or with a 'fibrous sheath' (principal piece), or with neither of these components (end-piece). Rat sperm tails contain no cytoplasmic or accessory tubules, so it is not possible to compare 9 + 2 tubules and cytoplasmic tubules within the same cells. It is possible, however, to compare 9 + 2 tubules in sperm tails with cytoplasmic tubules in nearby Sertoli cells, for the latter have larger numbers of microtubules in their cytoplasmic extensions (Christensen, 1965; Behnke, 1965 a). Pepsin digestion of sections of normal testes. When sections of rat testes were digested with pepsin, the first effect observed was disappearance of A-tubules, resulting in visible holes; in the same cells the central tubules and a segment of the B-tubule wall were not affected (Fig. 17). The holes seen after digestion of the A-tubules were always larger than the A-tubules themselves (Fig. 17). The next effect observed was the disappearance of the B-tubules and central tubules,. without leaving holes (Fig. 19). After extended digestion there was sometimes evidence of a badly denned 'hole' at the position of the central tubules (Fig. 19), but distinct circular holes such as occurred upon digestion of the A-tubules were never seen. Classes of microtubules 179

Table 1. Effects of experimental treatments on Nephrotoma suturalis microtubules

Spermatids A t 9 + 2 tubules Spermato- A \ cytes: Experimental Cytoplasmic Accessory Central spindle treatment microtubules tubules tubules B-tubules A-tubules tubules Glutaraldehyde fixation + + + + + + OsO4 fixation + or - + + + + o °C, is min - + + + + — 40 °C, 25 min + . + + + 5° °C 10 min - . + + + — 15 min, weak effect - . + or - + or - + — 15 min, medium effect - . - - + — 15 min, strong effect — . — — — — 1 "3 x io~4 M colchicine, 40 min — + + + + + or - Pepsin Weak effect + + + ± (part nor- — mal, part di- (holes) gested, leaving a hole) Strong effect _ _ _ — (one part — (no holes) (no holes) (no holes) holes) (holes) Negatively stained with - . + or - + (sometimes + phosphotungstate, pH 7 breaking up) + Indicates that microtubules were observed, as in normal cells, —, that no microtubules were observed. A point indicates that no observations were made.

In the mid-piece and principal piece the coarse fibres and fibrous sheath were also attacked by pepsin (Figs. 17-19). When a single section contained sperm tails sectioned at the level of the mid-piece and other sperm tails sectioned through the end-piece, it was seen that the tubules in the latter were more resistant to digestion than the corresponding tubules in the mid-piece. All tubules in the end-piece looked normal when the A-tubules in the mid- piece were digested (Fig. 17). Digestion of the A-tubules in the end-piece began when central and B-tubules had disappeared from the mid-piece. At this time, microtubules in adjacent Sertoli cells looked normal (Fig. 18). With prolonged digestion the B- tubules and central tubules disappeared from the end-piece (Fig. 19), and microtubules could not be seen in Sertoli cells. By comparing pepsin effects on tubules present in the same section, one can infer their relative sensitivities at the different levels of the sperm tail. These comparisons show that:

MD > Mc; MD > ED; Mc > Ec; PD > ED; and PD > MC)

where M is mid-piece, P is principal piece, E is end-piece, and subscripts D and c 180 O. Behnke and A. Forer refer to doublet and central tubules. We are not certain of the relationship between ED and Mc, nor between MD and PD. All these tubules, except perhaps the central tubules in the end-piece, were more sensitive than the cytoplasmic tubules in Sertoli cells. There was an overall loss of material from the control sections, but no differential loss of microtubules. Summarizing the results of pepsin digestion of rat sperm tails: the A-tubules and a part of the B-tubules were most sensitive, and the central tubules and a portion of the B-tubules less sensitive; the digestion of A-tubules resulted in holes, while the central tubules and portions of the B-tubules disappeared without leaving holes; finally, the 9 + 2 tubules in the mid-piece were more sensitive to pepsin digestion than were the corresponding tubules in the end-piece. Negatively stained rat sperm tubules. The rat sperm tails frayed only at the tip, and thus it was possible by this method to study only the tubules in that region. The main difference observed between rat sperm tubules and crane-fly spermatid tubules is that lateral projections were generally not seen in doublets from rat sperm (Fig. 29). Despite the lack of lateral projections, we can infer which is the A-tubule and which the B-tubule by comparison with the crane-fly spermatid doublets, where we saw lateral projections. The substructure of the tubules was similar to that of the crane-fly spermatid tubules. The individual tubules were composed of filaments about 30 A wide, with an interfilament distance (centre to centre) of about 60 A (Figs. 28, 29). We usually saw 5 filaments in A-tubules, 4 in B-tubules, and 6 in central tubules. Again, the total number of filaments per tubule is presumably about twice these values. Judged from the frequency with which they appeared intact, the central tubules were the least stable, the B-tubules were more stable, and the A-tubules were most stable. This is illustrated in Fig. 29. Summary. The experiments on rat sperm microtubules are summarized in Table 2. They show 3 of the same classes of tubules seen in crane-fly experiments, namely, A-tubules, B-tubules, and central tubules. Tubules in these 3 classes respond differently again from the cytoplasmic microtubules in Sertoli cells.

Cilia from rat trachea The experiments described above showed the similarities in microtubules from two quite different animals, crane fly and rat, but the tubules were in both cases from sperm- line cells. The experiments on cilia from rat trachea are more limited than those above, but illustrate the same general phenomena. Normal rat cilia contain the 9 + 2 tubules, as has been described for these and other cilia (Rhodin & Dalhamn, 1956). Pepsin digestion of sections of normal cilia. When sections of normal cilia were digested with pepsin, the first effect observed was the disappearance of both the A- tubules and a portion of the B-tubules, resulting in holes, while the central tubules looked normal. Then the central tubules and the remainder of the B-tubules disappeared. The central tubules left no holes. Sometimes there was a suggestion of poorly denned holes, or very small holes in the region of the two central tubules Classes of microtubules 181

Table 2. Effects of treatments on rat testis tubules

Sperm cells

Mid-piece End-piece

Central Central Sertoli Treatment tubules B-tubules A-tubules tubules B-tubules A-tubules cells Pepsin, + ± (part ab- - + + + + weak effect sent, with (holes) holes; part present.) Pepsin, — — (absent; - + ± (part ab- — + medium (absent, no part with (holes) sent, with (holes) effect holes) holes.) holes; part present.) Pepsin, — — — — — (absent; — — strong effect (no holes) (holes) (absent, no part with (holes) (no holes) holes) holes.) Negatively + + + stained with (sometimes (sometimes (always in- phosphotung- present) breaking up) tact) state, pH 5 + Indicates that microtubules were observed, as in normal cells, —, that no microtubules were observed. A point indicates that no observations were made.

(Fig. 20), but we never saw distinct circular holes such as were seen after digestion of the A-tubules. Negatively stained cilia. Cilia deprived of the surrounding membrane frayed into the 9 + 2 tubules, but the component microtubules remained joined at both the basal body (Fig. 30) and the tip, while spreading out in between. The doublet tubules had lateral projections (Fig. 31), which were found on the same tubule as those in crane- fly spermatid doublets. The tubules were composed of longitudinal filaments (Fig. 31), as in rat sperm and crane-fly spermatids.

DISCUSSION Classes of microtubules On the basis of their responses to the various experimental treatments, the tubules fall into 4 classes: (1) A-tubules; (2) B-tubules; (3) central tubules and accessory tubules; and (4) cytoplasmic tubules and spindle tubules (Tables 1,2). The members of each class responded in the same way to all experimental treatments, and each class of tubules responded differently from the other classes in at least one experimental treatment. The various kinds of tubules differ in position and surroundings. The A-tubules and B-tubules are joined into doublets, and each doublet is associated with one spoke. The central tubules are not joined as doublets, each is associated with 9 spokes, Table 3. Differences between 9 + 2 tubules

9+2 tubules 00 Cell Procedure A-tubules B-tubules Central tubules Remarks Reference Rooster (Gallus Shadow cast, after non-fixed Always present Always present Often not domesticus) sperm sperm were lysed in dis- present tilled water and dried down on a grid Grigg & Hodge, Shadow cast, after formalin- Present in mid- Present in mid- Not present 1940 fixed sperm were mixed piece piece with pepsin, fixed,an d dried down on a grid Various algae Shadow cast, after cells were Always present Always present Often not Fucus seratus Manton, 1953; fixed with osmium vapour present spermatozoid Manton, Clarke and sea water blotted & Greenwood, away 1953 Saprolegnia ferax, Manton, Clarke, and Chlorosaccus Greenwood & tdvaceus zoo- Flint, 1952 spores Sea urchin sperm Treated with distilled water, Not disrupted Not disrupted Easily dis- or sea water rupted Exact treatment Bradfield, 1955 Treated with distilled water, Not sensitive Not sensitive Sensitive ( not mentioned or digestive enzymes Trichonympha (a Negatively stained fiagella Always present Always present Rarely present Grimstone & a flagellate) obtained by lysing in i % Klug, 1966 phosphotungstate and One of the doublet tubules drying down on a grid collapses more than the other Tetrahymena (a Cilia isolated in TRIS-Mg"+. Present, always Present, but Not present Gibbons, 1963, ciliate) EDTA added. Study of sec- normal not infre- 196s tioned material quently distorted and/or r fragmented Diplodinium Sections of normal cells in Present Present Not present Occurs in the first Roth & Shigenaka, ecandatum (a ciliate) division. Osmium fixed stage of the nor- 1964 mally occurring resorption of cilia Rat sperm Sections of testes treated ex- Present. Cores Some present, Not present The first stage of perimentally by keeping are electron some not pre- sperm de- them in the abdominal dense sent generation cavity for varying times. Present. Elec- Not present Not present The second stage Nagano, 1963 Osmium fixed tron-dense of sperm de- cores generation Drosophila melano- Sections of normal testes. Electron-dense Hollow (not Electron-dense Daems, Persijn & gaster sperm (Methods not given) cores (grey) dense to elec- cores (black) Tates, 1963 trons) cores Sections of normal sperm, Cores contain Hollow cores Electron-dense The difference Acton, 1966 fixed with chilled osmium an electron- cores (grey) between A- and tetroxide dense body B-tubules was (black) not stated in the text, but was de- duced from Fig. 2 Table 3 {cont.)

Thermobia spermatids Sections of normal testes Not preserved Not preserved Preserveserved , and sperm fixed with KMnO 4 Bawa, 1964 Sections of normal testes Hollow cores Hollow cores Elecctron-densi e f fixed with osmium tetroxide ires > Acheta domestica Sections of normal testes Hollow cores Hollow cores; a Electron-dense Young sperma- spermatids and fixed with osmium tetroxide few have elec- cores tids sperm tron-dense cores Kaye, 1964 One of the doublet tubules has Electron-dense Mature sperm electron-dense (and structured) (and struc- cores tured) cores Tenebrio molitor Sections of normal testes Some have hoi- Hollow cores Electron-dense Cameron, 1965 mature sperm fixed with osmium tetroxide low cores, cores some have a dense body in their cores Macroglossum stella- Sections of normal testes Hollow cores Hollow cores Electron-dense Andre", 1961 tarum L. sperm fixed with osmium tetroxide cores Human sperm Sperm tails negatively stained Protofibrils not Protofibrils with phosphotungstate after readily seen; readily seen; The identifica- lysing in hypotonic saline somewhat not opaque tion of A-tubule and drying down opaque ap- and B-tubule Andre* & Thie'ry, pearance used by Andr6 1963 Oblique stria- Oblique stria- & Thie'ry is op- tions readily tions not posite to our seen readily seen identification Trichonympha Sections of normal ftagella, Occasionally Hollow cores Hollow cores Gibbons & Grim- 3 fixed with osmium tetroxide have electron- stone, i960 ©^ dense cores «• Rooster sperm Sections of normal testes, Electron-dense Hollow cores Hollow cores Nagano, i960, fixed either with osmium cores 1962 tetroxide, or acrolein followed by osmium tetroxide Fresh-water mussel Sections of normal gill fixed Electron-dense Hollow cores Hollow cores Gibbons, 1961 (Anodonta cataracta) with osmium tetroxide cores gills Various sperm tails Sections of tissue fixed with Electron-dense Hollow cores Hollow cores Lansing & Lamy, and cilia osmium tetroxide, or with cores i96i;Telkkae( glutaraldehyde followed by al. 1961; Nican- osmium tetroxide der & Bane, 1962; von Bons- dorff & Telkka, 1965; Fawcett & Ito, 1965

00 CO 184 O. Behnke and A. Forer and each is surrounded by the 9 doublets. The cytoplasmic microtubules are not joined as doublets, and they are neither associated with spokes nor regularly surrounded by other tubules. These differences in position and associated material could be the cause of the different responses to the experimental treatments. This does not seem to be the case, however, for the accessory tubules acted like the central tubules, yet they are outside the 9 doublets, and are not associated with the spokes. Furthermore, such a possibility does not explain the differences between the tubules after pepsin digestion, or heat treatment, or the differences in stability during negative- staining procedures (Tables 1, 2). Thus, the differences between the various tubules cannot be explained as being primarily due to differences in position and associated materials. The different responses to the experimental treatments might be due to different cellular responses, however, and might not represent actual physical or chemical differences in the tubules themselves. For example, we have found that cytoplasmic and spindle microtubules had disappeared after 15 min at o °C; this agrees with the observations of others on cytoplasmic microtubules (Tilney, cited by Tilney, Hiramoto & Marsland, 1966), and on spindle structure (Ris, 1949; Inou6, 1959, 1964; Harris & Bajer, 1965). When spindles are isolated from sea-urchin eggs, however, storage at 0 °C for prolonged periods does not cause spindle microtubules to disappear (Kane & Forer, 1965). Thus, spindle microtubules themselves are not affected by cold, and their disappearance when cells are placed at o °C must be a cellular response. This example might apply only to spindle microtubules, or to sea-urchin eggs, but nonetheless one must keep in mind the possibility that the various temperatures, osmium- tetroxide fixation, and colchicine treatments did not act directly on the tubules to produce the differences observed, but rather that the cell reacted to the treatment, and produced the different effects on the different tubules. In this respect, one should also leave open the possibility that o °C, 50 °C, and/or colchicine treatments did not cause microtubules actually to disappear, but rather that they changed the fixation properties of the microtubules, so that the latter were not preserved by the standard fixation, though actually present. There is evidence that this is indeed the case with microtubules of Actinosphaerium after pressure treatment (Tilney et al. 1966). How- ever, even if this is true, it doe9 not affect our general argument, for we observed differences between microtubules in one and the same cell. The differences in stability during negative staining, and the differences seen after pepsin digestion are difficult to explain as cellular responses, however, and suggest that the tubules are indeed physically and/or chemically different. We might conjecture from our results that although osmium tetroxide fixation is generally thought not to preserve microtubules, part of this might be due to the fact that the osmium tetroxide is often applied in the cold. Other workers have indicated that various kinds of microtubules in the 9 + 2 complex are different, and some such observations are summarized in Table 3. Each of these workers showed differences between two kinds of tubules, and, taken together, these observations illustrate differences between the various tubules which agree with those we have found. Classes ofmicrotubules 185 Our results on 9 + 2 tubules differ from those described by Grigg & Hodge (1949). We found the central tubules more resistant to pepsin than the 9 doublets, while Grigg & Hodge, working with fowl sperm, reported the reverse. They found that the entire tail was digested except for 9 tapering ' fibrils' in the mid-piece. The differences in results might be attributed to differences in species, or in technique, but it seems more likely that in the experiments of Grigg & Hodge pepsin digested the entire axial filament complex, in both mid-piece and end-piece, and that the 9 tapering remnant fibrils were the coarse bodies of the mid-piece. The effects of 50 °C on the 9 + 2 tubules of crane-fly spermatids greatly resemble the changes described in rat sperm tails by Nagano (1963), after rat testes had been placed in the abdominal cavity. This resemblance supports Nagano's supposition that the effects he observed were due to an increase in environmental temperature.

Differences along the length of microtubules The 9 + 2 tubules in the mid-piece of rat sperm tails were digested by pepsin more readily than the corresponding tubules in the end-piece. We assume that this is also true in crane-fly spermatids, where we cannot distinguish between middle- and end- piece, and we assume that this accounts for some of the variation observed after pepsin treatment of crane-fly spermatids. The differences between mid-piece and end-piece tubules might be due to inherent differences along the length of each of the tubules. It is reasonable to assume that the 9 + 2 tubules grow from one end, either from the tip or from the centriole. In that case one might expect a gradient of stability along the length of the tubules, and the differences in susceptibility (to pepsin) along the length of the tubules would reflect this. We think that this is the most likely explanation, but it is also possible that the difference along the length of the 9 + 2 tubules is not inherent in the tubules, but rather is due to other differences; for example, in fixation. If, for example, the fibrous sheath and coarse bodies acted as diffusion barriers and competitive binders of glutaraldehyde, it might be that the glutaraldehyde acted longer on the axial filament tubules in the end-piece than on the corresponding tubules in the mid-piece. Thus, the mid-piece tubules would probably be more easily digested than the corresponding end-piece tubules, because increased interaction of proteins with glutaraldehyde increases cross-linking and resistance to fragmentation (Quiocho & Richards, 1964). Our data indicate differences along the length of the 9 + 2 tubules. We have no similar data which apply to cytoplasmic or spindle microtubules. It is possible that there are such gradients in these other kinds of microtubules as well, and, in fact, there are data which indicate that this may be so in the axopods and cytosome of Actinosphaerium (Tilney & Porter, 1965; Tilney et al. 1966).

Substructure of microtubules Negative-staining experiments did not reveal consistent differences in substructure between the tubules, which were different by other criteria. The different kinds of microtubules were all composed of longitudinal filaments about 30 A wide, with an interfilament distance (centre to centre) of about 60 A. This substructure and these 186 O. Beknke and A. Forer dimensions are similar to those observed by others, on negatively stained axial filament, cytoplasmic, and spindle tubules from not closely related cells (e.g. Andre" & Thie'ry, 1963; Pease, 1963; Barnicot, 1966; Behnke & Zelander, 1966; Grimstone & Klug, 1966; Kiefer et al. 1966). The basic similarity in substructure between all these tubules argues that they are all constructed in basically the same way, and this makes even more puzzling our finding that there are distinct differences between the different kinds of tubules in one cell. Although we see no differences in the substructure of the negatively stained tubules, there might be differences which are not revealed in the negatively stained image. Grimstone & Klug (1966) used a combination of negative staining and optical diffrac- tion techniques and demonstrated that along the lengths of microtubules there were periodicities which were not readily seen by inspection of the micrographs. Perhaps the differences in stability which we have seen are due to differences in the periodic material which we do not see, but which were detected by Grimstone & Klug. Another possible source of the differences between the tubules might lie in the core material. That is, the tubule wall might be the same in all cases, but the tubules respond differently depending on the nature of the material within the core.

Composition of tubules Microtubules disappeared after digestion with pepsin. Thus they are composed of protein, at least in part. Since membranes were present when prolonged pepsin digestion had caused all microtubules to disappear, we must conclude that, contrary to the suggestion of others (Sandborn, Szeberenyi, Messier & Bois, 1965; Sandborn, 1966), the wall of the microtubules is different in composition from the membranes. Some tubules left holes after they were digested with pepsin, while others disappeared without leaving holes. We do not know what this difference is due to, but we can suggest some possibilities. In order for a structure to be attacked by pepsin, it must be in contact with the surface of the section, for pepsin does not seem to diffuse through Epon. Thus we presume that a structure will be completely digested only if the digestible material is topologically continuous with the surface. Since the microtubules invariably disappeared after digestion with pepsin, we conclude that the digestible tubule material is continuous throughout each tubule. To explain the disappearance of tubules without leaving holes, we presume that when tubule material is digested by pepsin, material remains which provides support for the non-digested material, so that no holes are formed and a general density remains. To explain the formation of holes, we presume that the pepsin digests some topologically continuous material, and the non-digested material which remains (if any) has no support, and disappears from the section during washing. For example, tubules which left holes and tubules which did not might be different in the packing of their component filaments ('protofibrils'). If the filaments did not actually touch but were separated from each other by a non-digestible 'matrix', the latter could give support after the filaments were digested, and thus no holes would be seen after digestion. If, on the other hand, the filaments were tightly packed, without much intermingling matrix, there would be Classes of microtubules 187 no supporting material remaining after the filaments were digested, and holes would be formed. Since there is loss of density from the entire A-tubule region in the early stages of formation of a hole, the possibility can be eliminated that only an outer shell is digested and the non-digested plug in the middle is washed away. This is not the only possible explanation, but it illustrates the kinds of restrictions which can be placed on possible explanations, and the kinds of reasoning which might lead to a satisfactory one. In this regard, we should point out that one need not restrict the considerations to differences between the walls of the tubules. For example, it is not unreasonable to suspect that some of the differences arise because different tubules have different material in their cores, for while some cytoplasmic microtubules appear hollow, others contain electron-dense 'dots' or other material (Gonatas & Robbins, 1964; Silveira & Porter, 1964; Bassot & Martoja, 1965, 1966; Behnke, 19656 and unpublished observations), and some 9 + 2 and accessory tubules regularly have an

(a) (c)

(f) (g) (0 Fig. 2. Interpretation of the structure of the A- and B-tubulea. (a) is a normal doublet. The B-tubule wall is composed of 2 different materials, represented as white (pepsin-resistant) and black areas. Both components are different from the material composing the A-tubule wall (shaded). (b)-{e) represent our interpretation of the various B-tubule abnormalities seen after storage at 50 °C (compare Figs. 8—11): the treatment causes separation of 'white', 'black' and 'grey' components of the doublets. (/MO represent our interpretation of the formation of the accessory tubules (compare Cameron, 1965): accessory tubules are formed from the pepsin-resistant (white) component of the B-tubules. electron-dense core (Gibbons, 1961; Telkka et al. 1961; Daems et al. 1963; Kaye, 1964; Cameron, 1965). It is also not unreasonable to suppose that the region immediately surrounding the tubules may be involved, for some microtubules have distinct clear areas around them (Ledbetter & Porter, 1963; Vivier & Schrevel, 1964; Behnke, 19656). Related to the problem of why some microtubules leave holes and others do not are the questions why one segment of the B-tubule was consistently more resistant than 188 O. Behnke and A. Forer the rest, and whether the resistant segment did or did not leave a hole when it finally disappeared. We assume that the B-tubule remnant did not leave a hole, and we suggest that each B-tubule may be composed of 2 kinds of material (Fig. 2 (a)), component I, adjacent to the A-tubule, and component II, the resistant segment, similar in composition to the central and accessory tubules. This assumption explains, first, why there is a more resistant segment of the B-tubule. Furthermore, heat treatment causes the B-tubule both to separate from the A-tubule and to fragment, and the remaining pieces could be interpreted as pieces of component I which had become separated from component II. This is illustrated diagrammatically in Fig. 2 (b)-(e) (compare .Figs. 8, 9). Similar configurations can be seen in Nagano (1963), Figs. 2, 7. Gibbons (1963, 1965) has shown that the arms can be separated chemically from the A-tubules of Tetrahymena cilia. When this was done, the A-tubules remained intact, but not uncommonly some of the B-tubules were fragmented and/or distorted. As- suming that the arms and component I have similar solubility properties, these dis- tortions can all be explained by the hypothesis that the B-tubules are composed of two components. Finally, Cameron (1965) has demonstrated that, in Tenebrio sperm, accessory tubules arise from the B-tubules, at a point on the B-tubules which corresponds, in our hypothesis, to the junction of the two components forming the B-tubule. By assuming that the accessory tubules are made from the pepsin-resistant component of the B-tubule, Fig. 2 (/)-(*)> one can explain why the accessory tubules in all cases acted like the central tubules (and the resistant segment of the B-tubule), as we have found (Tablesi, 2), and as has been seen in other cases (Kaye, 1964; Bawa, 1964; Cameron, 1965 Acton, 1966).

A. F. acknowledges the gracious hospitality of Dr E. Zeuthen, director of the Carlsberg Foundation Biological Institute, in the use of space, equipment and facilities. We acknowledge the support of the Danish State Research Fund (to O.B.) and the American Cancer Society (Postdoctoral Fellowship PF-249 to A.F.). We also acknowledge the assistance of Mr F. B. T. Birkes, without which this work could not have been completed.

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ABBREVIATIONS a A-tubule of 9 + 2 complex cf coarse fibres of rat sperm ac axial filament complex db dense body accessory microtubule er endoplasmic reticulum ar lateral projections from doublet tubules m mitochondrion b B-tubule of 9 + 2 complex mt cytoplasmic microtubule bb basal body P plasma membrane c central tubule t tonofilaments All micrographs of sectioned material are from tissue fixed with glutaraldehyde and post- fixed with osmium tetroxide.

Fig. 3. Part of a cross-sectioned spermatid tail from a crane fly. The axial filament complex (ac) consist of two central microtubules surrounded by 9 doublets. Nu- merous microtubules (mt) are seen in cross-section in the cytoplasm; some of these cytoplasmic microtubules tend to be grouped in pairs or in triplets (asterisks). The paired microtubules sometimes seem to be connected to each other by a fine line (arrows), x 100000. Fig. 4. Part of a crane-fly spermatid tail, more mature than that of Fig. 3, showing 2 axial filament complexes, both of which have accessory microtubules (am). Some cytoplasmic microtubules (mt) are seen around the mitochondrion (m). Two microtubules are apparently connected (arrow), x 94000. Fig. 5. Testis kept for 15 min at o°C in Ringer solution prior to fixation with glutaraldehyde. The section shows 4 axial filament complexes in one spermatid tail, all with accessory microtubules. Only 3 cytoplasmic microtubules are seen (arrows); the other cytoplasmic microtubules disappeared during the cold treatment (compare Fig. 4). The morphology of the 9 + 2 tubules and the accessory microtubules was not affected, x 52000. Journal of Cell Science, Vol. 2, No. 2 "Y

O. BEHNKE AND A. FORER (Facing p. 192) Fig. 6. Material treated in the same way as that in Fig. 5. No cytoplasmic microtubules are seen (compare Fig. 3). The 9 + 2 tubules seem unaffected, x 80000. Fig. 7. Testis kept in colchicine—Ringer solution for 60 min prior to fixation. The 9 + 2 tubules seem unaffected. No cytoplasmic microtubules can be seen, x 56000. Figs. 8-11. Testes kept for 15 min at 50 °C, in Ringer solution, prior to fixation with glutaraldehyde. Fig. 8. Section showing 2 axial filament complexes in one spermatid. The cytoplasmic microtubules have disappeared completely. One of the central tubules in each complex has also disappeared. Several of the B-tubules were damaged; some have lost their attachment to the A-tubules (asterisks), and others are fragmented (arrow). Some doublets appear normal, x 205000. Fig. 9. Section showing 2 axial filament complexes in one spermatid tail. The 2 central tubules appear intact, but some of the B-tubules of the peripheral doublets are detached from the A-tubules (arrows), x 126000. Fig. 10. The central and B-tubules have disappeared, and only the A-tubules are present. The remaining A-tubules are arranged roughly in a circle, x 108000. Fig. 11. The central and B-tubules have disappeared, and the remaining A-tubules are somewhat disordered, x 108000. Journal of Cell Science, Vol. 2, No. 2

O. REHNKE AND A. FORER Figs. 12-15. Micrographs showing the effect of pepsin digestion. Fig. 12. Crane-fly spermatid tails from a section treated with pepsin for 15 min. There are holes at the positions of the A-tubules; B-tubule remnants are discernible in some of the doublets (arrows). The central tubules, the accessory tubules, and the cytoplasmic tubules seem normal. Plasma membranes are intact, x 102000. Fig. 13. Crane-fly spermatid tail from a section treated with pepsin for 5 min. There are holes at the positions of the A-tubules; some B-tubule remnants can be seen. The cytoplasmic microtubules (arrows), central microtubules, and accessory microtubules seem normal, x 49000. Fig. 14. Crane-fly spermatid tail with 8 axial filament complexes, in a section treated with pepsin for 10 min. Holes can be seen at the sites of the doublets; central, accessory, and cytoplasmic microtubules seem normal, x 67000. Fig. 15. Crane-fly spermatid tail from a section treated with pepsin for 20 min. This shows an advanced stage of digestion. There are holes at the positions of the peripheral doublets. No B-tubule remnants can be seen. The central tubules cannot be seen, but there are no holes. Some cytoplasmic microtubules are barely discernible (arrows). The digested area immediately outside the axial filament complex is a 'dense body' (db) which is present in nearly mature sperm. The plasma membrane is intact, x 93000. Fig. 16. Two crane-fly spermatid tails from a section treated for 30 min with 01 N HC1. There is extraction of material from the section, but no differential loss of microtubules: 9 + 2, accessory, and cytoplasmic microtubules can be seen, x 42000. Journal of Cell Science, Vol. 2, No. 2

O. BEHNKE AND A. FORER Figs. 17-19. Pepsin treatment of Epon sections of normal rat sperm tails. Fig. 17. Transversely sectioned rat sperm tail principal pieces (upper), and an end- piece (lower left). The section was digested with pepsin for 10 min. In the principal pieces the A-tubules have been digested, leaving holes; portions of the B-tubules are visible (arrows), and the central tubules appear normal. All tubules in the end-piece seem normal. There is some digestion of the dense coarse fibres (cf). x 106000. Fig. 18. Principal pieces of two rat sperm tails, cut at different levels, one (left) with coarse fibres and the other (right) without coarse fibres. The section was treated with pepsin for 20 min. All 9 + 2 tubules are gone from the sperm tail to the left: the A-tubules left holes, the others did not. In the sperm tail to the right, the A-tubules left holes, while the central tubules and B-tubule remnants seem normal. The coarse fibres were also affected. The microtubules seen in a cytoplasmic extension of a Sertoli cell appear intact (arrow), x 59000. Fig. 19. A principal piece (right), and end pieces of rat sperm tails, in a section treated with pepsin for 15 min. In all of the sperm tails, the peripheral doublets have disappeared, leaving holes. The central tubules are not visible, but there are no holes. The outer coarse fibres and outer fibrous sheath of the principal piece have been digested, x 77000. Fig. 20. Cilia from rat trachea, sectioned transversely. The section was treated with pepsin for 20 min. The peripheral doublets have been digested, leaving holes. Central tubules are discernible in 2 cilia (arrows), x 52000. Fig. 21. Crane-fly spermarids ruptured on water, as in the negative-staining procedure, and then shadow-cast with platinum-carbon. The axial filament complex tubules remain as one group (ac). Numerous cytoplasmic microtubules are seen (nit) separate from the axial filament. Some cytoplasmic microtubules have broken into short fragments (arrow), x 30000. Journal of Cell Science, Vol. 2, No. 2

O. BEHNKE AND A. FORER Fig. 22. The 9 doublet tubules of the axial filament complex of a crane-fly spermatic! negatively stained with potassium phosphotungstate at pH 5. No central tubules are seen. In each doublet the A-tubules (a) are on the right and the B-tubules (6) are on the left. The A- and B-tubules are constructed of filaments, some of which are beaded (arrows), x 165000. Fig. 23. A crane-fly spermatid doublet tubule, negatively stained with potassium phosphotungstate at pH 7. The A-tubule shows projections (ar) arranged in pairs. x 165000. Fig. 24. Crane-fly spermatids ruptured on water, as in negative-staining experiments, and then shadow-cast with platinum-carbon. This figure shows doublet tubules from the 9 + 2 complex, and demonstrates the regularly spaced projections (arrows) found on one of the doublet tubules, x 36000. Fig. 25. Crane-fly spermatid 9 + 2 tubules negatively stained at pH 7. To the left are 4 doublets, and to the right are the 2 central tubules (c). The doublet and central tubules are constructed of filaments. The filamenti n one of the doublets shows distinct beading (arrow), x 207000. Journal of Cell Science, Vol. 2, No. 2

O. BEHNKE AND A. FORER Fig. 26. Crane-fly spermatid tubules negatively stained at pH 5. The axial filament complex (ac) and cytoplasmic microtubules (mt) can be seen. The cytoplasmic microtubules have a filamentous substructure similar to that of the axial filament complex tubules, x 160000. Fig. 27. Crane-fly spermatid cytoplasmic microtubules negatively stained at pH 5. x 235000. Fig. 28. Negatively stained preparation of rat sperm tail tips, stained with potassium phosphotungstate at pH 5. This micrograph shows 2 doublets, illustrating the filamentous substructure, x 200000. Journal of Cell Science, Vol. 2, No. 2

O. BEHNKE AND A. FORER Fig. 29. A preparation similar to that in Fig. 28, showing the 9 + 2 tubules. The central tubules (c) have frayed into their component filaments. The B-tubules indicated by asterisks also appear somewhat frayed, while all A-tubules appear intact, x 109000. Figs. 30, 31. Negatively stained preparations of the 9 + 2 complex of cilia from rat trachea. Fig. 30. Parts of the 9 + 2 complex of 3 cilia. Central tubules (c) are present. The doublets have lateral projections on one side, x 45000. Fig. 31. Higher magnification micrograph of 3 doublets, showing the filamentous substructure, x 175000. Journal of Cell Science, Vol. 2, No. 2

O. BEHNKE AND A. FORER