J. Cell Set. 51, 15-23 (1981) 15Printed in Great Britain © Company of Biologist! Limited ig8i

ACID PHOSPHATASE LOCALIZATION IN

PAS-BODIES OF GONYAULAX

RUTH E. SCHMITTER AND ANTONI J. JURKIEWICZBiology Department, University of Massachusetts at Boston, Boston, Mass. 02125,U.S.A.

SUMMARY

Periodic acid-Schiff staining, acid phosphatase localization, and yellow autofluorescencehave been correlated with the PAS-body of Gonyaulax polyedra for the first time. PAS-staining and acid phosphatase activity are both correlated with the PAS-body of Gonyaulaxtamarensis. These results suggest that the PAS-body of these marine dinoflagellate algaefunctions in subcellular digestion.

INTRODUCTION

It is important to know the subcellular localization of acid phosphatase activity inalgal cells. This enzyme is associated with digestive processes throughout the animalkingdom, but there are fewer reports of subcellular localizations from plant cells, andespecially from the algae. Many of the algal organisms studied were in unusualnutrient conditions or undergoing specific developmental changes. These include2 species of Euglena under conditions of carbon starvation and aging (Brandes, Buetow,Bertini & Malkoff, 1964; Malkoff & Buetow, 1964; Brandes, 1965; Sommer & Blum,1965; Palisano & Walne, 1972; Gomez, Harris & Walne, 19740,6); 3 species ofCryptomonas (Lucas, 1970); Polytomella caeca, a colourless heterotroph (Cooper,Bowen & Lloyd, 1974); Viva mutabilis during gamete release and fertilization (Briten,1975) and U.lactuca vegetative cells (Micalef, 1975); Micrasterias americana atvarious stages of growth (Noguchi, 1976); aging Ectocarpus sp. (Oliveira & Bisalputra,1977); and 2 wall-less volvocine algae, Dunaliellaprimolecta (Eyden, 1975) and Astero-monas gracilis (Swanson & Floyd, 1979).

Cultured Gonyaulax polyedra cells possess a large membrane-bounded sphericalbody, portions of which are stained by the periodic acid-Schiff (PAS) reaction.Schmitter (1971) proposed an active digestive function for these PAS-bodies, eitherin autophagy or in the use of stored metabolites, based in part on their ultrastructuralcontents: aggregates of electron-dense material, fibrous areas, and membranousvesicles. PAS-bodies are morphologically similar to the digestive granules in absorptivecells of Hydra (Slautterback, 1967) and to the food vacuoles of the dinoflagellatesCeratium hirundinella and Amyloodinium sp. (Dodge & Crawford, 1970; Lorn &Lawler, 1973). They are also similar in many respects to the Corps de Maupas ofCryptomonas reticulata (Lucas, 1970). They bear a lesser structural resemblance to the'accumulation bodies' of symbiotic dinoflagellates (Taylor, 1968, 1971a, b\ Tomas &

16 R. E. Schmitter and A. J. Jurkiewicz

Cox, 1973; Trench, 1974), in which granular electron-dense material increases withcell age until the contents often appear quite homogeneous in texture. Taylor (1968)suggested that the structure is an accumulation site for waste materials and Tomas &Cox (1973) proposed that the accumulation bodies of Peridinium balticum function inwaste storage or elimination. Lee (1977) has described accumulation bodies in free-living colourless heterotrophic isolates of Gyrodinium lebouriae Herdman but theseresemble PAS-bodies structurally more than accumulation bodies.

In our study we found acid phosphatase activity to be localized in the PAS-bodiesof the red tide organisms, Gonyaulax polyedra and G. tamarensis. This investigationalso supports the proposal of Schmitter (1971, 1973) that PAS-bodies of G. polyedrahave a digestive function.

MATERIALS AND METHODS

Growth of cells

G. polyedra and G. tamarensis were cultured in medium f/2 without silicate (Guillard &Ryther, 1962) at 21 and 16 °C, respectively, under an alternating cycle of 12 h light-12 h dark.G. polyedra strain GP60E was obtained from Dr R. R. L. Guillard, Woods Hole OceanographicInstitution; G. tamarensis was isolated by Dr C. Martin, University of Massachusetts MarineStation, Gloucester. All samples used were from cultures in the first hour of their light period(CT 0-1) unless otherwise stated.

Acid phosphatase localization

Cells were collected by centrifugation at 80-100g in an IEC clinical centrifuge for 4 min,then fixed with 2-5% glutaraldehyde in 005 M-sodium cacodylate buffer (pH 7-2), for 30 minat 3-4 °C. Subsequent steps were carried out in 12-ml centrifuge tubes; centrifugation be-tween steps was at 80-100g for 4 min. Acid phosphatase was localized within whole cells bya modification of Trelease's method (1980) for algae. Fixed cells were rinsed 3 times in caco-dylate buffer at room temperature before preincubating in 005 M-sodium acetate buffer(pH 5-o) for 20 min at room temperature. The acid phosphatase medium was mixed as followsto minimize precipitation: 0-36 M-lead nitrate was prepared using cooled, freshly boileddistilled water. One ml of lead nitrate solution was added in 20-fi\ aliquots with gentle sirringto n o ml of 0-05 M-sodium acetate buffer (pH 5-0) in which 0-3 g of sodium /?-glycerophosphatehad just been dissolved. The complete medium was preincubated at 37 °C for 1 h and anyprecipitate removed using Whatman no. 1 filter paper. Controls consisted of incubation mediumlacking substrate (Pb1+ control) and complete incubation medium containing the enzymeinhibitor sodium fluoride (NaF control). In the latter, sodium fluoride was added to the buffer-substrate mixture to give a final concentration of 001 M before lead nitrate was added. Controlsolutions were also preincubated and filtered. All incubations were carried out as 5 -ml volumesin stoppered centrifuge tubes for 30 min at 37 °C with occasional gentle stirring. The ensuingsteps were at room temperature. Cells were rinsed 3 times in distilled water, soaked for 5 minin i-o% acetic acid to remove unprecipitated lead, and rinsed again in distilled water. Sites oflead phosphate deposition were revealed by conversion to lead sulphide using i-o% aqueousammonium sulphide for 10 min. Cells were rinsed thoroughly with distilled water beforemounting in glycerol for light microscopy. Some preparations were stained with acetocarmine,so that nuclei were also clearly visible.

Cells were examined by light microscopy for acid phosphatase localization at a magnificationof x 400. Cells from several adjacent fields were scored for lead sulphide deposition withinPAS-bodiea; a minimum of 500 cells was counted from each slide. Photographs were recordedon Kodak Plus-X-Pan or Ektachrome 200 film.

Acid phosphatase in Gonyaulax 17

Periodic acid-Schiff staining

Periodic acid-Schiff (PAS) staining was carried out as described by Grimstone & Skaer (1972);their directions for Schiff's reagent were also used. Cells were fixed as for the acid phosphatasestudies, or at room temperature in ethanol/glacial acetic acid (3:1, v/v) for 30 min. Cells wererinsed thoroughly after either fixation. Non-specific staining of glutaraldehyde-fixed cells wasprevented by blocking with aniline/glacial acetic acid (1 :<), v/v) for 20 min. Pellets were gentlyresuspended throughout using a Pasteur pipette. Oxidation, staining, and rinsing times wereas described by Grimstone & Skaer, except that rinses were done by repeated centrifugation.Cells were mounted in glycerol and examined for PAS staining at a magnification of x 400.

Fluorescence microscopy

Autofluorescence of the PAS-bodies was studied using either unfixed cells, or cells fixed inthe ethanol/acetic acid fixative just described. Fixed cells were rinsed in distilled water twiceand mounted in glycerol for study with a Zeiss fluorescence microscope equipped with a high-intensity illuminator and superpressure mercury vapor lamp HBO 200 W/4. Excitation filtersBG3 and UGi were used in conjunction with a no. 53 barrier filter. Photographic records weremade using Kodak Technical Pan Film 2415 (ESTAR-AH base) and developed for 4 min withKodak D-19 developer for maximum contrast. At least 500 cells were examined at a magni-fication of x 400 in all studies. The autofluorescence is yellow.

RESULTS

Acid phosphatase

The highest levels of acid phosphatase activity were recorded when 2-5% glutar-aldehyde was employed. Lower concentrations of glutaraldehyde resulted in relativelylow levels of localization in identifiable PAS-bodies. For example, after 1*5%

Table 1. Acid phosphatase localization in Gonyaulax

Organism

G. polyedra

G. tamarensis

(days)

141729

3°352 1

% Acid phosphatase in PAS-bodies

Experimental

2423332445

13

Pb1+ control

0

0

0

0

0

0

NaF control

0

0

0

0

0

1

glutaraldehyde fixation a maximum of 9% of cells examined showed acid phosphataselocalization in PAS-bodies; Pb2+ controls gave up to 1% positive results. In ourhands methods other than that of Trelease (1980) resulted in a considerable degree oflead deposition within nuclei (e.g. see Gomori, 1952; Lewis & Knight, 1977). We didnot use any substrates other than /?-glycerophosphate in our studies (see Beaufay,1972).

Table 1 lists the results of several acid phosphatase localization experiments usingG. polyedra and one using G. tamarensis. The percentage of cells with lead sulphidedeposits in identifiable PAS-bodies is given for cultures of several ages. Figs. 4 and 7

R. E. Schmitter and A. J. Jurkicuricz

Acid phosphatase in Gonyaulax 19

show typical examples of localizations in G. polyedra; Fig. 3 gives a comparablepicture of G. tamarensis. Figs. 1 and 2 depict typical cells from NaF and Pb2+ controlsin G. tamarensis and G. polyedra, respectively.

The data do not allow any conclusions about the relative amounts of acid phos-phatase activity present in PAS-bodies of cells from cultures of different ages. Someof the reasons for this are discussed below.

A portion of cells in all experimental preparations stained non-specifically. We donot know whether this reflects intracellular disruption or some other undeterminedvariable. Such ' overstaining' was never observed in the NaF or Pb2+ controls.

Cells from the 29-day culture experiment were stained by the PAS reaction afteraniline blockage. A total of 35% of the experimental cells contained PAS-reactivePAS-bodies, as did 35% of the Pb2+ control cells.

The reasons for the lower percentage of localization in G. tamarensis are unknown.That experiment was done using the same incubation mixture as the 29-day G. poly-edra culture.

Fluorescence

Fluorescent PAS-bodies were seen in a high percentage of G.polyedra cells examinedfrom 3 times of day (14-day culture). Circadian times o and 7 h (light cycle), and16 h (dark cycle) had 90, 99, and 89% yellow autofluorescent PAS-bodies,respectively. Cells from o h showed no fluorescence at all if subjected to the PASreaction before viewing. A total of 98 % of these same cells had PAS-bodies whenviewed by conventional illumination. Fig. 5 depicts cells of G. polyedra as viewed byconventional illumination; Fig. 6 shows the same cells as viewed by fluorescencemicroscopy. The PAS-bodies are clearly visible as discrete sites of fluorescencesurrounded by indistinct cytoplasm.

Fig. 1. G. tamarensis; NaF control. Only 1 % of NaF control cells showed any leadsulphide deposition. Arrow indicates girdle region of cell. Clear area within is thenucleus. Figs. 1-5, bar 20/tm.Fig. 2. G. polyedra; Pbl+ control. None of the Pbl+ control cells showed lead sulphidedeposition in any of the experiments described.Fig. 3. G. tamarensis; acid phosphatase experimental preparation of 21-day culture.A total of 13 % of cells had lead sulphide localized in a PAS-body. Arrows indicatedeposits.Fig. 4. G. polyedra; acid phosphatase preparation of 35-day culture. A total of 45 %of cells had lead sulphide localized in PAS-bodies. Arrows indicate 3 deposits.Fig. 5. G. polyedra viewed by conventional illumination after fixation with ethanol/acetic acid.Fig. 6. Same cells as those of Fig. 5, viewed by fluorescence microscopy; 7-5 minphotographic exposure time. Discrete sites of fluorescence (PAS-bodies) are clearlyvisible, as are outlines of surrounding cytoplasm.

2O R. E. Schmitter and A. J. Jurkieioicz

er

Fig. 7. Two cells of G. polyedra with acid phosphatase localization in PAS-bodies.PAS-body in cell at right shows typical compact morphology. That at upper left hasseveral large granules at its periphery (arrows). Bar, 10 fim.Fig. 8. PAS-body of G. polyedra by electron microscopy. Methods were as given bySchmitter (1971). Two membrane-bound elements (1, 2) are closely associated withthe PAS-body, and another (3) is fusing with it. These elements may correspond tothe granular localizations noted in Fig. 7. ch, chloroplast; st, starch; er, endoplasmicreticulum. Bar, 1 fim.

Acid phosphatase in Gonyaulax 21

DISCUSSION

Results of the acid phosphatase studies show substantial localization within PAS-bodies. These results are not completely quantitative, however. First, it is known thatglutaraldehyde inhibits a portion of acid phosphatase activity, while at the same timepreserving sufficient activity for cytochemical localization (Sabatini, Bensch &Barrnett, 1963; Brunk & Ericsson, 1972). Swanson & Floyd (1979) determined fromin vitro studies of the /?-glycerophosphatase from Asteromonas that about 70% of theactivity of the enzyme was lost after the glutaraldehyde fixation method they employedfor cytochemistry. We have not done such a study, but the amounts of activitylocalizable in G. polyedra are comparable to the amount of activity remaining in theirstudy.

Second, we have not determined the pH optimum for acid phosphatase activity inG. polyedra. If it differs very much from pH 5-0, localizable activity would be lowerthan the optimum. Although we have not seen a study in which the pH optimum foracid phosphatase activity was much higher, Miiller (1973) has described acid phos-phatase activity from a trichomonad flagellate, with substantial activity above pH 6.

Third, because we used whole cells in our study, we cannot be certain that smalldeposits of lead sulphide would be visible by light microscopy. If elements such asthose associated with the PAS-body in Fig. 8 (labelled 1, 2, 3) contained activity andthe body-proper did not, we would not have been able to resolve them. Similarly, thecontents of PAS-bodies as seen by electron microscopy are not homogeneous. Acidphosphatase activity might be present only in certain portions at any given time.Small localizations would be visible only by electron microscopy. It is possible that thegranular deposits associated with the PAS-body in Fig. 7 do represent large mem-branous elements similar to those seen at the periphery of the PAS-body in Fig. 8.

The vegetative nucleus of G. polyedra is C-shaped, and the Golgi dictyosomes arearranged within the inner curve of the nucleus (Schmitter, 1971). Occasionally, weobserved granular deposits between the PAS-body and the inner curve of the nucleus.We are currently doing electron microscopic studies to investigate this point.

Correlations between autofluorescent cytoplasmic granules and lysosomal activityhave been made in a number of animal tissues (Koenig, 1963; Strehler, 1977). Pearse(1968) tentatively attributed this autofluorescence either to lipid or lipoprotein oflysosomal membranes, or to lipid material dissolved inside the organelles. Correlationshave also been made between lysosomal activity and PAS-reactive material (Koenig,1962; Strehler, 1977), and Koenig reported the disappearance of autofluorescence oflysosomes in certain tissues after the PAS-reaction. Studies on lipofuscin pigments inaging mammalian tissues (summarized by Strehler, 1977) have linked lysosomal PAS-reactivity and autofluorescence with the presence of partially oxidized, non-carbo-hydrate-containing lipids (Pearse, 1972). The fragments of membrane seen withinPAS-bodies in our study (Fig. 8; also by Schmitter, 1971) support the notion thatlipid composition of PAS-bodies is related to their autofluorescence and PAS-reactivity.

PAS-staining and acid phosphatase localization were observed by us in the same

22 R. E. Schmitter and A. J. Jurkievdcz

PAS-bodies of G. polyedra when cells were processed sequentially for acid phosphataselocalization and PAS-staining. Moreover, the size and unique location of PAS-bodieswithin G. polyedra make it possible to correlate PAS-staining, acid phosphataseactivity, and autofluorescence unequivocally. Gomez et al. (1974 a) unsuccessfullyattempted a similar correlation in aging Euglena gradlis. Their Euglena cells had beentreated with benzpyrene, so they were observing secondary lipid fluorescence, notautofluorescence (Pearse, 1968).

Our studies have shown that the PAS-bodies are a relatively permanent fixture inG. polyedra. Their acid phosphatase activity is probably, as originally proposed bySchmitter (1971), employed in autophagic processes recycling cellular materials. Theonly other alga so far reported to possess a permanent site for digestive processes, asopposed to the more usual transitory lysosomal activity, is Cryptomonas. Lucas (1970)localized acid phosphatase activity in the Corps de Maupas and has suggested that thestructure is permanent, since it can be observed in natural populations and culturesunder a variety of conditions. Lucas (1970) did not report any studies of PAS-stainingor autofluorescence of this structure.

Our study is the first report in which PAS-staining, acid phosphatase localization,and autofluorescence have all been correlated with the same subcellular algal organelle.

REFERENCES

BEAUFAY, H. (1972). The non-lysosomal localization of acid (p-nitro)-phenyl phosphataseactivity in various tissues. Appendix I to Methods for the Isolation of Lysosomes. In LysosomesA Laboratory Handbook (ed. J. T. Dingle), pp. 1-45. Amsterdam, London: North-Holland.

BRANDES, D. (1965). Observation on the apparent mode of formation of 'pure' lysosomes.J. Ultrastruct. Res. 12, 63-80.

BRANDES, D., BUETOW, D. E., BERTINI, F. & MALKOFF, D. B. (1964). Role of lysosomes incellular lyric processes. I. Effect of carbon starvation in Euglena gradlis. Expl molec. Path. 3,583-609.

BRATEN, T. (1975). Ultrastructure and localization of phosphohydrolases in gametes, zygotesand zoospores of Uha mutabilis Foyn. J. Cell Set. 17, 647-653.

BRUNK, U. T. & ERICSSON, J. L. E. (1972). The demonstration of acid phosphatase in in vitrocultured tissue cells. Studies on the significance of fixation, tonicity and permeability.Histochem. J. 4, 349-363.

COOPER, R. A., BOWEN, I. D. & LLOYD, D. (1974). The properties and subcellular localizationof acid phosphatases in the colorless alga, Polytomella caeca. J. Cell Sci. 15, 605-618.

DODGE, J. D. & CRAWFORD, R. M. (1970). The morphology and fine structure of Ceratiumhirundinella (Dinophyceae). J. Phycol. 6, 137-149.

EYDEN, B. P. (1975). Light and electron microscope study of Dunaliella primolecta Butcher(Volvocida). J. Protozool. 22, 336-344.

GOMEZ, M. P., HARRIS, J. B. & WALNE, P. L. (1974a). Studies of Euglena gradlis in agingcultures. I. Light microscopy and cytochemistry. Br. phycol. J. 9, 159—169.

GOMEZ, M. P., HARRIS, J. B. & WALNE, P. L. (1974ft). Studies of Euglena gradlis in agingcultures. II. Ultrastructure. Br. phycol. J. 9, 175-193.

GOMORI, G. (1952). Microscopic Histochemistry Prindples and Practice. Chicago: University ofChicago Press.

GRIMSTONE, A. V. & SKAER, R. J. (1972). A Guidebook to Microscopical Methods. Cambridge:Cambridge University Press.

GUILLARD, R. R. L. & RYTHER, J. (1962). Studies of marine planktonic diatoms. I. Cyclotellanana Hustedt and Detonula confervacea (Cleve) Gran. Can. J. Microbiol. 8, 229-239.

Acid phosphatase in Gonyaulax 23

KOENIG, H. (1962). Histological distribution of brain gangliosides: lysosomes as glycolipo-protein granules. Nature, Land. 195, 782-784.

KOENIG, H. (1963). The autofluorescence of lysosomes. Its value for the identification of lyso-somal constituents. .7. Hutochem. Cytochem. 11, 556—557.

LEE, R. E. (1977). Saprophytic and phagocytic isolates of the colourless heterotrophic dino-flagellate Gyrodinium lebouriae Herdman. J. mar. biol. Ass. U.K. 57, 303-315.

LEWIS, P. R. & KNIGHT, D. P. (1977). Staining Methods for Sectioned Material. In PracticalMethods in Electron Microscopy, vol. 5, part 1 (ed. A. M. Glauert). Amsterdam, New York,Oxford: North-Holland.

LOM, J. & LAWLOR, A. R. (1973). An ultrastructural study of the mode of attachment in dino-flagellates invading gills of Cyprinodontidae. Protistologica 9, 293-309.

LUCAS, I. A. N. (1970). Observations on the fine structure of the Cryptophyceae. I. The genusCryptomonas. J. Phycol. 6, 30-38.

MALKOFF, D. B. & BUETOW, D. E. (1964). Ultrastructure changes during carbon starvation inEuglena gracilis. Expl Cell Res. 35, 58-68.

MICALEF, H. (1975). Donnees compl6mentaires sur les caracteres cytomorphologiques etcytochimiques de la zone golgienne des cellules vegetatives de I'Ulva lactuca L. (Chloro-phycees, Ulvales). C. r. hebd. Sianc. Acad. Set., Paris D 281, 775-777.

MOLLER, M. (1973). Biochemical cytology of trichomonad flagellates. I. Subcellular localizationof hydrolases, dehydrogenases and catalase in Tritrichotnonas foetus. J. Cell Biol. 57, 453-474.

NOGUCHI, T. (1976). Phosphatase activities and osmium reduction in cell organelles of Micra-sterias americana. Protoplasma 87, 163-178.

OLIVEIRA, L. & BISALPUTRA, T. (1977). Ultrastructural and cytochemical studies on the natureand origin of the cytoplasmic inclusions of aging cells of Ectocarpus (Phaeophyta, Ecto-carpales). Phycologia 16, 235-243.

PALISANO, J. R. & WALNE, P. L. (1972). Acid phosphatase activity and ultrastructure of agedcells of Euglena granulata. J. Phycol. 8, 81-88.

PEARSE, A. G. E. (1968). Histochemistry, Theoretical and Applied, vol. 1, 3rd edn. Edinburgh,London: Churchill Livingstone.

PEARSE, A. G. E. (1972). Histochemistry, Theoretical and Applied, vol. 2, 3rd edn. Edinburgh,London: Churchill Livingstone.

SABATINI, D. D., BENSCH, K. & BARRNETT, R. J. (1963). Cytochemistry and electron micro-scopy. The preservation of cellular ultrastructure and enzymatic activity by aldehydefixation. J. Cell Biol. 17, 19-58.

SCHMITTER, R. E. (1971). The fine structure of Gonyaulax polyedra, a bioluminescent dino-flagellate. J. Cell Sci. 9, 147-173.

SCHMITTER, R. E. (1973). Structural and functional aspects of the paniculate bioluminescenceof Gonyaulax polyedra. Ph.D. thesis, Harvard University.

SLAUTTERBACK, D. B. (1967). Coated vesicles in absorptive cells of Hydra. J. Cell Sci. 2,563-572-

SOMMER, J. R. & BLUM, J. J. (1965). Cytochemical localization of acid phosphatases in Euglenagracilis. J. Cell Biol. 24, 235-251.

STREHLER, B. L. (1977). Time, Cells, and Aging, 2nd edn. New York, San Francisco, London:Academic Press.

SWANSON, J. & FLOYD, G. L. (1979). Acid phosphatase in Asteromonas gracilis (Chlorophyceae,Volvocales): a biochemical and cytochemical characterization. Phycologia 18, 362-368.

TAYLOR, D. L. (1968). In situ studies on the cytochemistry and ultrastructure of a symbioticdinoflagellate. J. mar. biol. Ass. U.K. 48, 348-366.

TAYLOR, D. L. (1971a). Ultrastructure of the ' zooxanthella' Endodinium chattonii Hovassein situ. J. mar. biol. Ass. U.K. 51, 227-234.

TAYLOR, D. L. (19716). On the symbiosis between Amphidinium klebsii (Dinophyceae) andAmphiscolops langerhansi (Turbellaria: Acoela). J. mar. biol. Ass. U.K. 51, 301-313.

TOMAS, R. N. & Cox, E. R. (1973). Observations on the symbiosis of Peridinium balticum andits intracellular alga. I. Ultrastructure. J. Phycol. 9, 304-323.

TRELEASE, R. N. (1980). Cytochemical localization. In Handbook of PhycologicalMethods, vol. 3,Developmental and Cytological Methods (ed. E. Gantt), pp. 305-318. Cambridge, London,New York, New Rochelle, Melbourne, Sydney: Cambridge University Press.

TRENCH, R. K. (1974). Nutritional potentials in Zoanthus sociathus (Coelenterata, Anthozoa).Helgola'nder wiss. Meeresunters. 26, 174-216.

(Received 18 March 1981)