Preliminary ? 6..
3. 4. 5. 6. 7. 8. 9. 10. II. 12, 13. 14.
Miller, D A, Rev, V G, Tantravahi, R & Miller, 0 J, Exp cell res 101 (1976) 235. Miller, 0 J, Miller, D A, Dev, V G. Tantravahi, R & Croce, C M, Proc natl acad sci US 73 (1976) 4531. Croce, C M, Talavera, A, Basilica, C & Miller, 0 J. Proc natl acad sci US 74 (1977) 694. Elideiri, G L, J cell biol53 (19?2) 1?7. Toniclo, D & Basilica, C, Nature 248 (1974) 411. Miller, 0 J, Dev, V G, Miller, D A, Tantravahi, R & Eliceiri, G L, Exp cell res 115 (1978) 457. Davidson, R & Gerald, P, Som cell gen 2 (1976) 165. Littlefield, J, Proc natl acad sci US 50 (1963) 568. Miller, 0 J, Miller, D A, Kouri, R E. Allderdice, P W, Dev, V G, Grewal, M S & Hutton, J J, Proc natl acad sci US 68 (1971) 1530. Goodpasture, C & Bloom, S E, Chromosoma 53 (1975) 37. Dev, V G, Miller, D A & Rechsteiner, M, J cell biol79 (1978) 353a. Dev, V, Tantravahi, R, Miller, D & Miller, 0, Genetics 86 (1977) 389. Horak, I, Coon, H & Dawid, I, Proc natl acad sci US 71 (1974) 1828.
Received May 14, 1979 Revised version received July 27, 1979 Accepted July 30, 1979
Printed in Sweden Copyright 0 1979 by Academic Press. Inc. All rights of ~p~uction in any form reserved 0014-4827/79/120429-06$02.00/O
The intracellular location of adenylyl cyclase in the cellular slime molds ~i~~yos~eliurn dis~o~eum and Polysphondylium pallidum RUDOLF HINTERMANN and ROGER W. PARISH, Plant Biology Institute, University of Zikich, CH-8008
Zzirich,
Switzerland
notes
429
in the absence of nutrients, single cells of Dictyostelium discoideum begin to secrete cyclic AMP (CAMP). This induces neighbouring cells both to respond chemotactitally and themselves synthesize and release CAMP [l-3]. This CAMP signalling leads to the formation of multicellular aggregates which subsequently develop into fruiting bodies [4]. The CAMP binds to receptors on the ceil surface [5, 61. However, the means by which adenylyl cyclase activity is stimulated and endogenous CAMP released are not known, although cGMP may be involved [7, 81. Cytochemistry indicates that adenylyl cyclase is located on the cytoplasmic surface of the plasma membrane [9]. Maeda & Gerisch [lo] have indirect ultrastructural evidence that CAMP is accumulated in vesicles and vacuoles which then fuse with the plasma membrane, releasing CAMP to the extracellular space. In such a model the location of adenylyl cyclase in the vesicle membrane could be postulated, intravesicular or cytoplasmic ATP serving as substrate [lo]. Plasma membrane location of the cyclase would require rapid transport of newly synthesized CAMP into the vesicles. In this paper we have used cell fractionation techniques to examine the intracellular location of adenylyl cyclase. Materials
and Methods
discoideum NC-4 (wild type) cells were grown in liquid medium with E. colt’ [I l] and Polysphondylium pallidurn (strain Ti-1) cells with E. coli on agar [12]. Cells were washed with 15 mM phosphate buffer (pH 6.8). Plasma membranes were isolated using the concanavalin A-Triton X-100 method [13]. Phagocytic vacuoles were isolated by flotation in sucrose gradients after the cells had phagocytosed latex beads [14]. Nuclei were isolated according to Charlesworth & Parish [ 1.51. Cells were homogenized in SF medium [16] (0.5 M sorbitol, 2.5% Ficoll, 2 mM ditbiothreitol, 1 mM EDTA in 0.05 M Tris-HCl, pH 7.4). Homogenization (4°C) was carried out using a Potter homogenizer with a Teflon pestle attached to an electric drill. The cell concentration was 10” ml-’ and after 50-70 strokes
Dictyosteliutn
Summary. Adenylyl cyclase activity was low or not detectable on intact cells and in isolated plasma membranes, phagocytic vacuoles and nuclei of the two slime mold species examined. The entire activity of homogenates was sedimentable and concentrated in a light membrane fraction. When this fraction was centrifugated through sucrose density gradients the adenylyl cyclase activity sedimented differently from all other enzymes measured. The gradient fractions with the highest specific activity of adenylyl cyclase consisted mainly of small vesicles. No changes in adenylyl cyclase distribution were associated with development. The possibility that cellular slime mold adenylyl cyclase activity is associated with vesicles in vivo, as already suggested by Maeda & Gerisch [lo], is discussed. 28-791813
Exp Ct4Rr.s
123 (1979)
430
Prel~mi~a~
Table 1. Adenylyl (Ti-1) cells
notes
cyclase in subcellularfractions
Fraction
Protein in the assay (l-4
Whole cells (NC-4 and Ti-I) Homogenate Ti-1
(10’ cells ml-‘) 2oOXlO
Homogenate NC-4
200-400
150000 S (Ti-1)
X-100
Plasma membrane Ti- 1
20-60
NC-4 Nuclei (vegetative cells) Ti-1 NC-4 Phagocytic vacuoles NC-4
20-60
of
D. discoideum
(NC-4)
and P. pullidum
Adenylyl cyclase activity (pmoles CAMP mg protein-’ min-‘) n.d. 0.71 (vegetative cells) 1.85 (aggregating cells) 0.3 (vegetative cells) 8.0 (aggregating cells) 1.09 (vegetative cells) 6.14 (aggregating cells) n.d. (vegetative cells) 0.22 (starved cells) nd.
38 63 347
0.22 0.36 0.03
68
n.d.
n.d., not detectable. whole cells were centrifugated out, resuspended in SF medium, further homogenized (50-70 strokes) and intact cells again removed. The supematants were combined (‘homogenate’) and subjected to differential centrifugation. Four fractions were isolated: 35 (3 Ooo g,,, 20 minf, 30s (30~ g,,, 20 min), 1.50s (150~ g,,, 60 min) and supematant. Homogenates were centrifuged at 10000 g,, for 20 min, the supernatant removed and centrifuged at 150000 g,, for 60 min. The sediment was resuspended in SF medium using a Kontes glass homogenizer (4°C). The suspension was layered onto linear 4 ml sucrose gradients @o-45% w/v) over a 0.3 ml 60% (w/v) sucrose cushion in 2 mM dithiothreitol, 1 mM EDTA and 0.05 M Tris-HCl buffer, pH 7.8. The gradients were centrifnged for 2 h at 190000 gaV using a Spinco L65b ultracentrifuge. Activities of alkaline phosphatase, acid phosphatase, malate dehydrogenase, a-mannoside, /3-N-acetylglucosaminidase and NADPH-dichlorophenolindopheno1 oxidoreductase were determined as previously described [14, 17, 181. Adenylyl cyclase was measured according to Salomon et al. [19] (‘modified method c’). CAMP phosphodiesterase, ATPase and AMPase activities were assayed using **C-1abelled substrates and separating products with thin-layer chromatography. Preparation and examination of gradient fractions with the electron microscope were performed as described [20].
Results and Discassio~
The adenylyl cyclase activity in homogenates of D. discoideum and P. pallidum ExpCeilRes
123 (1979/
cells depended on the developments stage at which cells were harvested (table 1) (cf [21, 221). We isolated various subcellular fractions and measured the specific activity of adenylyl cyclase. The majority (more than 90%) of activity was sedimentable, suggesting the enzyme was associated with a membrane fraction. (Inhibitors were not detected in the supematant.) In agreement with other results [21], no activity was found associated with the cell surface (table 1). Plasma membranes generally lacked adenylyl cyclase activity; although low levels were occasionally detectable they were less than the specific activity of the original homogenate (table 1). However, the Triton X-100 used in the plasma membrane isolation may have partly inhibited the enzyme. Famham [9] has presented cytochemical evidence that adenylyl cyclase is associated with the cytoplasmic surface of the plasma membrane. However, we found phagocytic vacuoles contained no or barely detectable activity (max. 4 cpm above a background of
Preliminary
notes
43 1
80 70 60 50 40 30
1
I
20 IO 6 0
I ”
12
3
4
150 s
Fig. 1. Abscissa: the three sedimentable fractions; ordinate: % of sedimentable spec. act. Distribution of enzyme activities between the three membrane fractions isolated by differential centrifugation (see Methods). (I) NADPH oxido-reductase; (2) malate dehydrogenase; (3) o-mannosidase; (4) /3-
N-acetylglucosaminidase; (5) alkaline phosphatase; and (6) adenylyl cyclase. P. pallidurn is shown; similar distributions were obtained using D. discoideum ceils. No changes in distribution were found between different developmental stages.
20 cpm) (table 1). This result argues against a plasma membrane location of the enzyme unless regions of the membrane containing adenylyl cyclase are not phagocytosed. Cytochemical studies carried out in our laboratory (U. Mtiller, unpublished) have been unable to confirm the results of Farnham [9]. Some adenylyl cyclase activity was consistently found associated with nuclei (table 1). However, the activity was variable and relatively low and thus may have been due to contamination. Differential centrifugation indicated adenylyl cyclase activity was largely associated with a light (slowly sedimenting) membrane fraction (fig. 1). Enzymes known to be present in the plasma membrane (alkaline phosphatase [ 141, CAMP phosphodiesterase [23], Mg2+-ATPase (Parish & Weibe1 [27]), 5’-AMPase [24]), mitochondrial marker enzymes and acid hydrolases associated with vacuoles ([14], Parish & Muller [28]) were differently distributed among the fractions (fig. 1).
When a light membrane fraction was centrifuged through a density gradient (20-45 % sucrose w/v), we consistently found the highest activities of adenylyl cyclase in the middle regions of the gradient (e.g. density 1.123 g cmp3) (fig. 2). Marker enzymes for vacuoles and plasma membranes were most active in the denser regions of the gradient (fig. 2). The activities of 5’-AMPase, Mg2+ATPase and CAMP phosphodiesterase resembled alkaline phosphatase in their distribution (not shown). Electron microscopy showed small vesicles in the adenylyl cyclase enriched fractions from the middle of the gradient (fig. 3a). The majority of these vesicles were less than 0.2 pm in diameter, the size chosen by Maeda & Gerisch [lo] to distinguish between vesicles and vacuoles. They found the temporal increase of small vesicles and vacuoles closely resembled that of intracellular CAMP, both during spontaneous oscillations and following stimulation by an extracellular pulse [lo]. We also found similar vesicles in the denser fractions from the Err, Cd Res 123(1979~
~ b -
i
1 I-1
IL
[
1
i 5-s
s -13
Fig. 2. Abscissa: (a) fraction no.; (b) combined fractions from gradient (a); ordinate: (a) (tefi) alkaline and acid phosphatase activities (A&,X 103x60 min-’ in 25 ~1 samples), protein (425 ~1); (right) adenylyl cyclase act. (pmoles cAMPx 10 mitt-‘/25 ~1); (b) spec. act. (a) Distribution of enzyme activitiesin a 2045 % (w/v) sucrose density gradient followjng ~ent~fugation of the fraction sedimenting between 10000 g (20 mitt) and 150000 g (60 min). A, Acid phosphatase; D, alkaline phosphatase; A, protein; 0, adenylyl cyclase.
gradient, along with larger vesicles (‘vacuoles’), some mitochondria and large membrane fragments (fig. 3b). Thus the distribution of small vesicles does parallel the adenylyl cyclase distribution in gradients. However, the possibility that the vesicles are homogenization artifacts cannot be dismissed, and final proof that such adenylyl cyclase containing vesicles exist in vivo will depend on the development of reliable cytochemical methods. The vesicles postulated to release CAMP from DictyosteIium cells may be comparExp Ceil Res 123 (1979)
(b) Specific activity of three enzymes in three regions of the gradient. Fractions from the lower, middle and upper regions of the gradient were combined, diluted, centrifuged (150000 g, 60 min) and enzyme activities in the sedimented membrane determined. The enzymes are: (1) acid phosphatase; (2) alkaline phosphatase (A~~xmg protein’x30 min’); (3) adenylyl cyclase (pmoies cAMPxmg protein’xmin-‘f. D. discoideum amoebae were used; similar results were obtained with P. pallidurn.
able to the presynaptic vesicles involved in transmitter storage and release from cholinergic synapses [lo]. Nevertheless, two points must be taken into consideration. (1) The plasma membrane contains at best low adenylyl cyclase activity. So, if fusion between the postulated vesicles and the plasma membrane does occur, removal of Fig. 3. Eiectronmi~rographs of two fractions from the gradient depicted in tig. 2. (a) Middle (fraction 7); (b) lower (fraction 3) region of the gradient. X22000. Bar, 1 pm.
Preliminary
notes
433
434
Preliminary
notes
the vesicle membrane must be rapid or the adenylyl cyclase activity be inhibited. (2) The localization of the adenylyl cyclase activity was similar in P. pallidurn and D. discoideum, although the former does not utilize CAMP as its acrasin [25]. There is, however, evidence that CAMP does function chemotactically in the migrating grex of P. pallidum and may be involved in its development [26]. The localization of adenylyl cyclase in a specific light membrane (eventually small vesicle) fraction may be related to the control of enzyme activity. This work was supported by the Schweizerischer Nationalfonds zur FGrderung der wissenschaftlichen Forschung (grant no. 3.673-0.75).
References 1. Konijn, T M, Adv cyclic nucleo res 1 (1972) 17. 2. Gerisch, G & Malchow, D, Adv cyclic nucleo res 7 (1976) 49. 3. Schaffer, B M, Nature 255 (1975) 549. 4. Bonner, J T, The cellular slime molds. Princeton Universitv Press. Princeton (19671. 5. Hendersdn, E J, j biol chem‘250 (1975) 4130. 6. Green. A A & Newell. PC. Cell 6 (1975) 129. 7. Wurster, B, Schubiger’, U, Wick, u‘ & derisch, G, FEBS lett 76 (1977) 141. 8. Mata, J M, I&ens; F A, van Haastert, P J M & Konijn, TM, Proc natl acad sci US 74 (1977) 2348. 9. Famham, C J M, Exp cell res 91 (1975) 36. 10. Maeda, Y & Gerisch, G, Exp cell res 110 (1977) 119. 11. Sussman, M, Methods in cell physiology (ed D Prescott) vol. 2, p. 397. Academic Press, New York (1966). 12. Raper, K B, Quart rev bio126 (1951) 169. 13. Parish, R W & Miiller, U, FEBS lett 63 (1976) 40. 14. Parish. R W & Pelli. C. FEBS lett 48 (1974) 293. 15. Charlesworth, M C’& Parish, R W, Eur j biochem 75 (1977) 241. 16. - Ibid 54 (1975) 307. 17. Parish, R W, Eur j biochem 58 (1975) 523. 18. - Planta 123 (1975) 15. 19. Salomon, Y, Londos, C & Rodbell, M, Anal biochem 58 (1974) 541. 20. Parish, R W; Eurj biochem 22 (1971) 423. 21. Klein, C, FEBS lett 68 (1976) 125. 22. - FEMS lett 1 (1977) 17. 23. Malchow, D, Nggele, B, Schwarz, H & Gerisch. G, Eur j biochem 28 (1972) 136. 24. Green, A A & Newell, P C, Biochem j 140 (1974) 313. 25. Bonner, J T, Hall, E M, Noller, S, Oleson, F B & Roberts, A B, Devl bio129 (1972) 402. 26. Jones, M E & Robertson, A, J cell sci 22 (1976) 41.
27. Parish, R W & Weibel, M. Submitted for publication. 28. Parish, R W & Miiller, U. In preparation. Received May 22, 1979 Accepted July 11, 1979
Printed in Sweden Copyright 0 1979 by Academic Press, Inc. All rights of reproduction in any form reserved 0014-4827/79/120434-08$0?.00/0
The hyaline layer is a collagen-containing extracellular matrix in sea urchin embryos and reaggregating cells EVELYN
SPIEGEL
and MELVIN
SPIEGEL,
partment of Biological Sciences, Dartmouth Hanover, NH 03755, and Marine Biological tory, Woods Hole, MA 02543, USA
DeCollege, Labora-
Evidence is given to support the classification of the hyaline layer of sea urchin embryos and reaggregating cells as a collagen-containing extracellular matrix. Ruthenium red staining shows the presence of striated fibril-like structures, dense spheroids, crystalline lattice structures and filamentous material. Collagenase digestion causes disappearance of the fibril-like structures; hyaluronidase treatment causes a diminution of other matrix components, but does not affect the fibrillar structures. The hyaline layer maintains the integrity of the embryo and is also involved in morphogenesis, which are among the functions of extracellular matrices. Summary.
The hyaline layer (HL) of sea urchin embryos plays an important role in early development. The firm attachment of the cells to the HL maintains the shape of the embryo and the orientation of the cells during cleavage. During invagination, the integrity of the ectoderm depends largely on the HL, which maintains the cohesion and orientation of the cells as the ectodermal layer moves in and the embryo changes shape [l, 4,
101.
When the HL is removed by treating embryos with calcium-free sea water, the cells lose their adhesion to each other and the embryos fall apart [ 131. These dissociated cells are able to form lamellipodia, move about and reaggregate to form almost nor-
3. 4. 5. 6. 7. 8. 9. 10. II. 12, 13. 14.
Miller, D A, Rev, V G, Tantravahi, R & Miller, 0 J, Exp cell res 101 (1976) 235. Miller, 0 J, Miller, D A, Dev, V G. Tantravahi, R & Croce, C M, Proc natl acad sci US 73 (1976) 4531. Croce, C M, Talavera, A, Basilica, C & Miller, 0 J. Proc natl acad sci US 74 (1977) 694. Elideiri, G L, J cell biol53 (19?2) 1?7. Toniclo, D & Basilica, C, Nature 248 (1974) 411. Miller, 0 J, Dev, V G, Miller, D A, Tantravahi, R & Eliceiri, G L, Exp cell res 115 (1978) 457. Davidson, R & Gerald, P, Som cell gen 2 (1976) 165. Littlefield, J, Proc natl acad sci US 50 (1963) 568. Miller, 0 J, Miller, D A, Kouri, R E. Allderdice, P W, Dev, V G, Grewal, M S & Hutton, J J, Proc natl acad sci US 68 (1971) 1530. Goodpasture, C & Bloom, S E, Chromosoma 53 (1975) 37. Dev, V G, Miller, D A & Rechsteiner, M, J cell biol79 (1978) 353a. Dev, V, Tantravahi, R, Miller, D & Miller, 0, Genetics 86 (1977) 389. Horak, I, Coon, H & Dawid, I, Proc natl acad sci US 71 (1974) 1828.
Received May 14, 1979 Revised version received July 27, 1979 Accepted July 30, 1979
Printed in Sweden Copyright 0 1979 by Academic Press. Inc. All rights of ~p~uction in any form reserved 0014-4827/79/120429-06$02.00/O
The intracellular location of adenylyl cyclase in the cellular slime molds ~i~~yos~eliurn dis~o~eum and Polysphondylium pallidum RUDOLF HINTERMANN and ROGER W. PARISH, Plant Biology Institute, University of Zikich, CH-8008
Zzirich,
Switzerland
notes
429
in the absence of nutrients, single cells of Dictyostelium discoideum begin to secrete cyclic AMP (CAMP). This induces neighbouring cells both to respond chemotactitally and themselves synthesize and release CAMP [l-3]. This CAMP signalling leads to the formation of multicellular aggregates which subsequently develop into fruiting bodies [4]. The CAMP binds to receptors on the ceil surface [5, 61. However, the means by which adenylyl cyclase activity is stimulated and endogenous CAMP released are not known, although cGMP may be involved [7, 81. Cytochemistry indicates that adenylyl cyclase is located on the cytoplasmic surface of the plasma membrane [9]. Maeda & Gerisch [lo] have indirect ultrastructural evidence that CAMP is accumulated in vesicles and vacuoles which then fuse with the plasma membrane, releasing CAMP to the extracellular space. In such a model the location of adenylyl cyclase in the vesicle membrane could be postulated, intravesicular or cytoplasmic ATP serving as substrate [lo]. Plasma membrane location of the cyclase would require rapid transport of newly synthesized CAMP into the vesicles. In this paper we have used cell fractionation techniques to examine the intracellular location of adenylyl cyclase. Materials
and Methods
discoideum NC-4 (wild type) cells were grown in liquid medium with E. colt’ [I l] and Polysphondylium pallidurn (strain Ti-1) cells with E. coli on agar [12]. Cells were washed with 15 mM phosphate buffer (pH 6.8). Plasma membranes were isolated using the concanavalin A-Triton X-100 method [13]. Phagocytic vacuoles were isolated by flotation in sucrose gradients after the cells had phagocytosed latex beads [14]. Nuclei were isolated according to Charlesworth & Parish [ 1.51. Cells were homogenized in SF medium [16] (0.5 M sorbitol, 2.5% Ficoll, 2 mM ditbiothreitol, 1 mM EDTA in 0.05 M Tris-HCl, pH 7.4). Homogenization (4°C) was carried out using a Potter homogenizer with a Teflon pestle attached to an electric drill. The cell concentration was 10” ml-’ and after 50-70 strokes
Dictyosteliutn
Summary. Adenylyl cyclase activity was low or not detectable on intact cells and in isolated plasma membranes, phagocytic vacuoles and nuclei of the two slime mold species examined. The entire activity of homogenates was sedimentable and concentrated in a light membrane fraction. When this fraction was centrifugated through sucrose density gradients the adenylyl cyclase activity sedimented differently from all other enzymes measured. The gradient fractions with the highest specific activity of adenylyl cyclase consisted mainly of small vesicles. No changes in adenylyl cyclase distribution were associated with development. The possibility that cellular slime mold adenylyl cyclase activity is associated with vesicles in vivo, as already suggested by Maeda & Gerisch [lo], is discussed. 28-791813
Exp Ct4Rr.s
123 (1979)
430
Prel~mi~a~
Table 1. Adenylyl (Ti-1) cells
notes
cyclase in subcellularfractions
Fraction
Protein in the assay (l-4
Whole cells (NC-4 and Ti-I) Homogenate Ti-1
(10’ cells ml-‘) 2oOXlO
Homogenate NC-4
200-400
150000 S (Ti-1)
X-100
Plasma membrane Ti- 1
20-60
NC-4 Nuclei (vegetative cells) Ti-1 NC-4 Phagocytic vacuoles NC-4
20-60
of
D. discoideum
(NC-4)
and P. pullidum
Adenylyl cyclase activity (pmoles CAMP mg protein-’ min-‘) n.d. 0.71 (vegetative cells) 1.85 (aggregating cells) 0.3 (vegetative cells) 8.0 (aggregating cells) 1.09 (vegetative cells) 6.14 (aggregating cells) n.d. (vegetative cells) 0.22 (starved cells) nd.
38 63 347
0.22 0.36 0.03
68
n.d.
n.d., not detectable. whole cells were centrifugated out, resuspended in SF medium, further homogenized (50-70 strokes) and intact cells again removed. The supematants were combined (‘homogenate’) and subjected to differential centrifugation. Four fractions were isolated: 35 (3 Ooo g,,, 20 minf, 30s (30~ g,,, 20 min), 1.50s (150~ g,,, 60 min) and supematant. Homogenates were centrifuged at 10000 g,, for 20 min, the supernatant removed and centrifuged at 150000 g,, for 60 min. The sediment was resuspended in SF medium using a Kontes glass homogenizer (4°C). The suspension was layered onto linear 4 ml sucrose gradients @o-45% w/v) over a 0.3 ml 60% (w/v) sucrose cushion in 2 mM dithiothreitol, 1 mM EDTA and 0.05 M Tris-HCl buffer, pH 7.8. The gradients were centrifnged for 2 h at 190000 gaV using a Spinco L65b ultracentrifuge. Activities of alkaline phosphatase, acid phosphatase, malate dehydrogenase, a-mannoside, /3-N-acetylglucosaminidase and NADPH-dichlorophenolindopheno1 oxidoreductase were determined as previously described [14, 17, 181. Adenylyl cyclase was measured according to Salomon et al. [19] (‘modified method c’). CAMP phosphodiesterase, ATPase and AMPase activities were assayed using **C-1abelled substrates and separating products with thin-layer chromatography. Preparation and examination of gradient fractions with the electron microscope were performed as described [20].
Results and Discassio~
The adenylyl cyclase activity in homogenates of D. discoideum and P. pallidum ExpCeilRes
123 (1979/
cells depended on the developments stage at which cells were harvested (table 1) (cf [21, 221). We isolated various subcellular fractions and measured the specific activity of adenylyl cyclase. The majority (more than 90%) of activity was sedimentable, suggesting the enzyme was associated with a membrane fraction. (Inhibitors were not detected in the supematant.) In agreement with other results [21], no activity was found associated with the cell surface (table 1). Plasma membranes generally lacked adenylyl cyclase activity; although low levels were occasionally detectable they were less than the specific activity of the original homogenate (table 1). However, the Triton X-100 used in the plasma membrane isolation may have partly inhibited the enzyme. Famham [9] has presented cytochemical evidence that adenylyl cyclase is associated with the cytoplasmic surface of the plasma membrane. However, we found phagocytic vacuoles contained no or barely detectable activity (max. 4 cpm above a background of
Preliminary
notes
43 1
80 70 60 50 40 30
1
I
20 IO 6 0
I ”
12
3
4
150 s
Fig. 1. Abscissa: the three sedimentable fractions; ordinate: % of sedimentable spec. act. Distribution of enzyme activities between the three membrane fractions isolated by differential centrifugation (see Methods). (I) NADPH oxido-reductase; (2) malate dehydrogenase; (3) o-mannosidase; (4) /3-
N-acetylglucosaminidase; (5) alkaline phosphatase; and (6) adenylyl cyclase. P. pallidurn is shown; similar distributions were obtained using D. discoideum ceils. No changes in distribution were found between different developmental stages.
20 cpm) (table 1). This result argues against a plasma membrane location of the enzyme unless regions of the membrane containing adenylyl cyclase are not phagocytosed. Cytochemical studies carried out in our laboratory (U. Mtiller, unpublished) have been unable to confirm the results of Farnham [9]. Some adenylyl cyclase activity was consistently found associated with nuclei (table 1). However, the activity was variable and relatively low and thus may have been due to contamination. Differential centrifugation indicated adenylyl cyclase activity was largely associated with a light (slowly sedimenting) membrane fraction (fig. 1). Enzymes known to be present in the plasma membrane (alkaline phosphatase [ 141, CAMP phosphodiesterase [23], Mg2+-ATPase (Parish & Weibe1 [27]), 5’-AMPase [24]), mitochondrial marker enzymes and acid hydrolases associated with vacuoles ([14], Parish & Muller [28]) were differently distributed among the fractions (fig. 1).
When a light membrane fraction was centrifuged through a density gradient (20-45 % sucrose w/v), we consistently found the highest activities of adenylyl cyclase in the middle regions of the gradient (e.g. density 1.123 g cmp3) (fig. 2). Marker enzymes for vacuoles and plasma membranes were most active in the denser regions of the gradient (fig. 2). The activities of 5’-AMPase, Mg2+ATPase and CAMP phosphodiesterase resembled alkaline phosphatase in their distribution (not shown). Electron microscopy showed small vesicles in the adenylyl cyclase enriched fractions from the middle of the gradient (fig. 3a). The majority of these vesicles were less than 0.2 pm in diameter, the size chosen by Maeda & Gerisch [lo] to distinguish between vesicles and vacuoles. They found the temporal increase of small vesicles and vacuoles closely resembled that of intracellular CAMP, both during spontaneous oscillations and following stimulation by an extracellular pulse [lo]. We also found similar vesicles in the denser fractions from the Err, Cd Res 123(1979~
~ b -
i
1 I-1
IL
[
1
i 5-s
s -13
Fig. 2. Abscissa: (a) fraction no.; (b) combined fractions from gradient (a); ordinate: (a) (tefi) alkaline and acid phosphatase activities (A&,X 103x60 min-’ in 25 ~1 samples), protein (425 ~1); (right) adenylyl cyclase act. (pmoles cAMPx 10 mitt-‘/25 ~1); (b) spec. act. (a) Distribution of enzyme activitiesin a 2045 % (w/v) sucrose density gradient followjng ~ent~fugation of the fraction sedimenting between 10000 g (20 mitt) and 150000 g (60 min). A, Acid phosphatase; D, alkaline phosphatase; A, protein; 0, adenylyl cyclase.
gradient, along with larger vesicles (‘vacuoles’), some mitochondria and large membrane fragments (fig. 3b). Thus the distribution of small vesicles does parallel the adenylyl cyclase distribution in gradients. However, the possibility that the vesicles are homogenization artifacts cannot be dismissed, and final proof that such adenylyl cyclase containing vesicles exist in vivo will depend on the development of reliable cytochemical methods. The vesicles postulated to release CAMP from DictyosteIium cells may be comparExp Ceil Res 123 (1979)
(b) Specific activity of three enzymes in three regions of the gradient. Fractions from the lower, middle and upper regions of the gradient were combined, diluted, centrifuged (150000 g, 60 min) and enzyme activities in the sedimented membrane determined. The enzymes are: (1) acid phosphatase; (2) alkaline phosphatase (A~~xmg protein’x30 min’); (3) adenylyl cyclase (pmoies cAMPxmg protein’xmin-‘f. D. discoideum amoebae were used; similar results were obtained with P. pallidurn.
able to the presynaptic vesicles involved in transmitter storage and release from cholinergic synapses [lo]. Nevertheless, two points must be taken into consideration. (1) The plasma membrane contains at best low adenylyl cyclase activity. So, if fusion between the postulated vesicles and the plasma membrane does occur, removal of Fig. 3. Eiectronmi~rographs of two fractions from the gradient depicted in tig. 2. (a) Middle (fraction 7); (b) lower (fraction 3) region of the gradient. X22000. Bar, 1 pm.
Preliminary
notes
433
434
Preliminary
notes
the vesicle membrane must be rapid or the adenylyl cyclase activity be inhibited. (2) The localization of the adenylyl cyclase activity was similar in P. pallidurn and D. discoideum, although the former does not utilize CAMP as its acrasin [25]. There is, however, evidence that CAMP does function chemotactically in the migrating grex of P. pallidum and may be involved in its development [26]. The localization of adenylyl cyclase in a specific light membrane (eventually small vesicle) fraction may be related to the control of enzyme activity. This work was supported by the Schweizerischer Nationalfonds zur FGrderung der wissenschaftlichen Forschung (grant no. 3.673-0.75).
References 1. Konijn, T M, Adv cyclic nucleo res 1 (1972) 17. 2. Gerisch, G & Malchow, D, Adv cyclic nucleo res 7 (1976) 49. 3. Schaffer, B M, Nature 255 (1975) 549. 4. Bonner, J T, The cellular slime molds. Princeton Universitv Press. Princeton (19671. 5. Hendersdn, E J, j biol chem‘250 (1975) 4130. 6. Green. A A & Newell. PC. Cell 6 (1975) 129. 7. Wurster, B, Schubiger’, U, Wick, u‘ & derisch, G, FEBS lett 76 (1977) 141. 8. Mata, J M, I&ens; F A, van Haastert, P J M & Konijn, TM, Proc natl acad sci US 74 (1977) 2348. 9. Famham, C J M, Exp cell res 91 (1975) 36. 10. Maeda, Y & Gerisch, G, Exp cell res 110 (1977) 119. 11. Sussman, M, Methods in cell physiology (ed D Prescott) vol. 2, p. 397. Academic Press, New York (1966). 12. Raper, K B, Quart rev bio126 (1951) 169. 13. Parish, R W & Miiller, U, FEBS lett 63 (1976) 40. 14. Parish. R W & Pelli. C. FEBS lett 48 (1974) 293. 15. Charlesworth, M C’& Parish, R W, Eur j biochem 75 (1977) 241. 16. - Ibid 54 (1975) 307. 17. Parish, R W, Eur j biochem 58 (1975) 523. 18. - Planta 123 (1975) 15. 19. Salomon, Y, Londos, C & Rodbell, M, Anal biochem 58 (1974) 541. 20. Parish, R W; Eurj biochem 22 (1971) 423. 21. Klein, C, FEBS lett 68 (1976) 125. 22. - FEMS lett 1 (1977) 17. 23. Malchow, D, Nggele, B, Schwarz, H & Gerisch. G, Eur j biochem 28 (1972) 136. 24. Green, A A & Newell, P C, Biochem j 140 (1974) 313. 25. Bonner, J T, Hall, E M, Noller, S, Oleson, F B & Roberts, A B, Devl bio129 (1972) 402. 26. Jones, M E & Robertson, A, J cell sci 22 (1976) 41.
27. Parish, R W & Weibel, M. Submitted for publication. 28. Parish, R W & Miiller, U. In preparation. Received May 22, 1979 Accepted July 11, 1979
Printed in Sweden Copyright 0 1979 by Academic Press, Inc. All rights of reproduction in any form reserved 0014-4827/79/120434-08$0?.00/0
The hyaline layer is a collagen-containing extracellular matrix in sea urchin embryos and reaggregating cells EVELYN
SPIEGEL
and MELVIN
SPIEGEL,
partment of Biological Sciences, Dartmouth Hanover, NH 03755, and Marine Biological tory, Woods Hole, MA 02543, USA
DeCollege, Labora-
Evidence is given to support the classification of the hyaline layer of sea urchin embryos and reaggregating cells as a collagen-containing extracellular matrix. Ruthenium red staining shows the presence of striated fibril-like structures, dense spheroids, crystalline lattice structures and filamentous material. Collagenase digestion causes disappearance of the fibril-like structures; hyaluronidase treatment causes a diminution of other matrix components, but does not affect the fibrillar structures. The hyaline layer maintains the integrity of the embryo and is also involved in morphogenesis, which are among the functions of extracellular matrices. Summary.
The hyaline layer (HL) of sea urchin embryos plays an important role in early development. The firm attachment of the cells to the HL maintains the shape of the embryo and the orientation of the cells during cleavage. During invagination, the integrity of the ectoderm depends largely on the HL, which maintains the cohesion and orientation of the cells as the ectodermal layer moves in and the embryo changes shape [l, 4,
101.
When the HL is removed by treating embryos with calcium-free sea water, the cells lose their adhesion to each other and the embryos fall apart [ 131. These dissociated cells are able to form lamellipodia, move about and reaggregate to form almost nor-
