Morphological Studies on the Transcription of Spinach Chloroplast DNA

Morphological Studies on the Transcription of Spinach Chloroplast DNA R. J. ROSE and A. G. C. LINDBECK Department of Biological Sciences, The University of Newcastle, N .S.W., 2308 Australia Received September 18, 1981 . Accepted February 27,1982

Summary Morphological observations have been made on spinach chloro~last DNA transcription. Vesicles obtained from isolated chloroplasts treated with 3.5 mM Mg + and whole chloroplasts lysed with sodium-N-lauroyl sarcosinate have been used. Both preparations were collected on grids, stained with phosphotungstic acid and uranyl acetate, and viewed in the electron microscope. Structural networks are associated with the vesicles, and are interpreted as thylakoid-associated DNA involved in transcription. Ribosome-size granules are associated with the structural network. Detergent-lysed chloroplasts reveal strucures consisting of DNA filaments with lateral transcription complexes. Granules are associated with the complexes and their possible nature is discussed. The transcriptional complexes, besides occurring in association with the thylakoids, would also contribute to the morphology of the stromal matrix.

Key words: Spinacia oleracea L., chloroplast, chloroplast DNA, transcription.

Introduction Chloroplasts of higher plants contain many copies of chloroplast DNA (Herrmann and Possingham, 1980); recently in young mesophyll cells it has been estimated that there are 200-300cpDNA molecules per chloroplast (Lampp a and Bendich, 1979; Scott and Possingham, 1980). In many species the cpDNA copies are spread throughout the chloroplast in many nucleoid areas (Kowallik and Herrmann, 1972; Rose and Possingham, 1976; Kuroiwa et al., 1981). In spinach there is evidence from light microscope autoradiography (Possingham and Rose, 1976), electron microscope autoradiography (Rose and Possingham, 1976) and DAPI staining (Scott and Possingham, 1980) for the nucleoid areas being spread throughout the chloroplast. However, in wheat leaves DAPI staining reveals that cpDNA is located at the periphery of the chloroplast (Sellden and Leech, 1981). The number of nucleoid areas and the number of cpDNA copies depends on the stage of development of the chloroplast (Kowallik and Herrmann, 1974; Lamppa et al., 1980; Scott and Possingham, 1980). The cpDNA can be membrane-associated (Woodcock and Fernandez-Moran, 1968; Herrmann et al., 1974; Rose and Possingham, 1976) and is probably in the form of a Abbreviations: cpDNA, chloroplast DNA; cpm, counts per minute; DAP!, 4'6-diamidino-2phenylindole; RNP, ribonucleoprotein; sarkosyl, sodium-N-lauroyl sarcosinate.

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folded chromosome (Herrmann et al., 1974; Turischeva, 1975; Yoshida et al., 1980). There is evidence in spinach (Rose, 1979) and in pea chloroplasts (Filippovich et al., 1975) that the membrane association of cpDNA is mainly with granal thylakoids. There is little morphological information on how the approximately 45 J.1m (Herrmann and Possingham, 1980) circular cpDNA molecules are organized in situ for transcription. We have obtained information on this, using vesicles obtained from isolated chloroplasts treated with 3.5 mM Mg2+ and whole chloroplasts lysed with sarkosyl.

Materials and Methods Vesicle Preparations Photosynthetic membrane vesicles were prepared by isolating and preparing chloroplasts from near fully grown spinach leaves as previously described (Rose, 1979), washing and resuspending the chloroplasts in 3.5 mM Mg2+ (as MgCh), and storing overnight at 4°C. The vesicles were fixed in 4 % formaldehyde, dried on to carbon-coated copper grids (200 mesh) and stained with phosphotungstic acid (1 % in 50% ethanol) for one minute. After rinsing in 50% ethanol, the vesicles were stained with uranyl acetate (1 % in 50 % ethanol) for one minute. The grids were rinsed in 50 %, 70 %, 95 % and 100 % ethanol and air dried prior to examining in a lEOL 100CX electron microscope. Vesicles labelled with 3H-thymidine or 3H-uridine were obtained from cultured spinach disks as previously described (Rose, 1979). About 30 disks per plate were incubated in 1.85 MBq (6)H) thymidine (740GBq/mM) or 1.85MBq (5_ 3H) uridine (1.11 TBq/mM), added in 1 ml to the surface of the 20 ml of nutrient agar medium in 9 cm Petri dishes. The radiochemicals were from the Radiochemical Centre, Amersham, U.K. Equal aliquots of labelled vesicles were treated for 3 h with 0.1 % Calbiochem DNAase II in 3 mM MgS04 , pH 6.5 at 37° (Lord and Lafontaine, 1969),0.2 % Sigma RNAase A at pH 6.8 at 37° (Lord and Lafontaine, 1969), or left untreated at 37°. At the conclusion of the incubation the vesicles were collected as a 1,000g pellet, washed in 3.5 mM Mg2+ and counted in a toluene-based scintillation fluid.

Lysed Chloroplasts Chloroplasts from 50 mm spinach leaves were isolated in 0.4 M sucrose plus 3.5 mM Mg2+ and separated from nuclei as previously described (Rose, 1979), removed from the 20 %/60 % sucrose gradient interface and aliquots (15 Ilg chlorophyll) treated with an equal volume of 4 % sarkosyl. Immediately on lysis the preparation was centrifuged at 5,000 rpm for 5 minutes on to a carbon-coated grid through a 25 x 4 mm cushion of 0.1 M sucrose plus 10% formaldehyde, in a perspex device which fitted into the swinging bucket of a Sorvall HB-4 rotor. Grids were rinsed in 0.4 % Kodak Photo-Flo, dried, then stained as for the vesicle preparation, and rinsed in 50 %, 70 %, 95 %, 100 % ethanol and isopentane and air dried (Miller et aI., 1970). In some preparations more intact chloroplasts were obtained using chloroplasts isolated in the sorbitolbased Buffer A as described by Frankel et a1. (1979).

Results Chloroplast preparations isolated free of nuclei and treated with 3.5 mM Mg2+ lose their outer envelope membranes and form vesicles bounded by swollen stroma lamellae membranes, with grana which are more resistant to swelling on the surface of the vesicles (Rose, 1979). These vesicles when dried on to grids and stained with Z. Pjlanzenphysiol. Bd. 106. S. 129-137. 1982.

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Fig. 1: Vesicle formed by treating chloroplasts in 3.5 mM Mg2+. Network of fibres with associated granules (arrowhead) associated with areas where there a number of membrane layers. Bar is 1 J.1m. Fig. 2: A network of fibres with associated granules in association with another vesicle. Bar is 0.5 J.1m. Inset shows discrete granules (arrowheads). Bar is 100 nm. Fig. 3: A network similar to that in Fig. 2 after DNAase II treatment prior to staining. Bar is 1 J.1m. Inset shows a branched fibre, with granules (arrowheads). Bar is 200 nm.

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phosphotungstic acid and uranyl acetate have on their surface (Fig. 1) a network of fibres with associated granules (Fig. 2). The individual granules have diameter (measured as a median diameter if ovoid in shape) of about 19 nm (19.2±0.5 nm, Fig. 2 inset). The dark band on the vesicle is probably a granal area, suggestive of stacking (Fig. 1). It is these latter areas which usually have the previously described networks associated with them. Previous autoradiographic studies (Rose, 1979) have shown the DNA of the vesicles associated with the grana, as have our unpublished data with the fluorochrome DAPI which stains cpDNA (Coleman, 1978). The morphological observations would suggest a relationship between the granules and the membrane bound cpDNA. Studies on this latter point were made by treating vesicles on grids with DNAase II for I hour prior to staining. The network of fibres with the associated granules fragmented following the DNAase treatment (Fig. 3), suggesting that the integrity of the network is DNA dependent. Some of the structures remaining showed a resemblance to transcription complexes (Miller and Hamkalo, 1972), in the sense that there is a central fibre with lateral fibres and granules (Fig. 3 and Fig. 3 inset), which prompted subsequent morphological studies using a Miller-type method. The relationship between DNA, RNA and the vesicle was further examined by obtaining vesicles labelled with 3H-thymidine or 3H-uridine and treating the vesicle with nucleases (Table 1). DNAase removed both labelled DNA and RNA, while RNAase only removed labelled RNA from the vesicle. These data can be explained if the cpDNA is membrane bound and is associated with RNA transcripts. Table 1: Effect of nucleases on labelled vesicles. Treatment 3H-Thymidine 3H-Uridine labelled vesicles labelled vesicles (cpm)") (cpm)*) Control 34,683 RNAase A 27,129 DNAase II 7,424

29,624 1,779 5,967

*) Radioactivity determined as counts per minute per aliquot using liquid scintillation spectrometry.

To obtain further morphological information on the putative transcription complexes, attempts were made to spread out the complexes by using procedures similar to those developed by Miller et al. (1970). Chloroplast preparations free of nuclei were lysed with sarkosyl and centrifuged on to carbon-coated grids. Results of such treatment are shown in Figs. 4 -7. Long single fibres are clearly visible, and granular aggregates occur as lateral appendages to these fibres (Fig. 4). When granules that are clearly single are measured, they are about 24 nm (23.5 ±0.5 nm) in diameter while the central fibres measure about 3 nm (2.9 ± 0.3 nm) in diameter (Figs. 4 -7). In situations where granules are spread out more, they appear to have an association with a Z. P/lanzenphysiol. Bd. 106. S. 129-137. 1982.

Fig.4: Low magnification view of cpDNA fibres (larger arrowhead) and associated granular aggregates (smaller arrowhead). Bar is 111m. Fig. 5: Individual cpDNA fibre (largest arrowhead) with associated transcriptional complexes. Bar is 0.5 11m. Inset shows transcriptional complex. Medium-sized arrowhead points to a fine fibril and small arrowheads indicate a row granules. Bar is 100 nm. Fig. 6: A transcriptional complex (smaller arrowhead indicates a single granule) is attached to a DNA strand (larger arrowhead) via a light staining granule. Bar is 100 nm. Fig.7: Individual cpDNA fibre (larger arrowhead) with associated transcriptional complexes (smaller arrowhead). Bar is 0.5 11m. Inset shows transcriptional complexes. Largest arrowhead is DNA. Intermediate-sized arrowhead indicates a fine fibril and small arrowhead indicates single granules. Bar is 100 nm.

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fine fibril (Fig. 5 inset, Fig. 7 inset). Treatment of grids with DNAase for 3 h prior to staining removed the long 3 nm strands and granular aggregates were left on the grid, confirming that the 3 nm strands were DNA. However, RNAase treatment for 3 h did not break up the granular aggregates. We have found it difficult to visualize neatly growing transcripts, i.e. those showing a progressive increase in length. This could be related to the rate of RNA initiation. However, in more intact chloroplasts than those used in Figs. 4 -7, we have observed what appears to be lengths of DNA with more transcription, but material (presumably protein and lipid) coating the whole structure (including DNA) gave less clarity to the transcriptional complexes. The light staining granule in Fig. 6 warrants comment, as it is in the expected position for RNA polymerase, but appears to be too large to be the molecule. However, isolated RNA polymerase, DNA complexes of bacteria show the molecule to be very irregular in appearance and to approach the size of ribosomes (Lubin, 1969). At least some areas of the genome showed gene segments which were associated with very fine lateral fibrils, and these would most likely be ribosomal genes. DNA fibres showing no transcription activity were also present. Discussion

The vesicles with attached DNA used in this study have been previously described (Rose, 1979). As shown by light microscopy and thin sections examined in the electron microscope, they consist of swollen stroma membranes with attached grana on their surface. Areas with a number of membrane layers on the dried down vesicle (Fig. 1) would correspond to grana, and it is these areas which usually have the structures seen in Figs. 1-3. The structural networks associated with the vesicle are interpreted as thylakoidassociated DNA involved in transcription. The evidence for this is that DNA and RNA are associated with the vesicle (Table 1) and the complex is dependent on DNA for its integrity (Table 1, Fig. 3). The granule size (Fig. 2, inset) is similar to that of chloroplast ribosomes (Gunning and Steer, 1975). Extensive data are available showing polyribosome associations with chloroplast thylakoids (Chua et aI., 1973; Margulies and Michaels, 1975; Alscher et aI., 1978; Margulies and Weistrop, 1980). It is possible then that if nascent mRNA is present, there is a close association between transcription and translation, however, the evidence is circumstantial. The nature of the vesicle preparation is such that the transcriptional complexes are aggregated when dried on to a grid. When the Miller technique is employed the transcriptional complexes are spread out and show the relationship of transcription to the DNA fibre more clearly. The 3 nm diameter obtained for the DNA fibres compares with the 2.5 nm found in sectioned chloroplasts by Ris and Plaut (1962) and the 4 nm found by Miller et al. (1970) for Escherichia coli active gene preparations. The diameter of double helix DNA is about 2 nm (Miller et aI., 1970). Z. Pjlanzenphysiol. Bd. 106. S. 129-137. 1982.

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The 24 nm granules in the lateral structures are larger than those found in the dried down vesicles (19 nm). These structures could, however, be ribosomes thickened due to protein or lipid coating the granules during lysis. RNAase insensitivity of the transcript is probably because of the inability of the enzyme to gain access to mRNA. Some isolated chloroplast polyribosomes obtained by detergent lysis of membranes are RNAase resistant (Margulies and Michaels, 1975). Those granules shown in the insets of Figs. 5 and 7 where an associated fine fibril is just visible are about 20 nm in diameter. Chloroplast ribosomes usually have a diameter of 17 - 20 nm in thin sections (Gunning and Steer, 1975) or 22 x 17 nm in purified negatively stained preparations (Bruskov and Odinstova, 1968). The transcription complexes shown in Figs. 5, 6 and 7 do resemble those observed in bacteria (Miller et aI., 1970) and mitochondria (Laird et aI., 1973) where coupled translation and transcription occur. Preliminary studies in osmotically shocked Euglena chloroplasts have also suggested coupled transcription and translation in chloroplasts (Miller and Hamkalo, 1972). Gibbs et aI. (1974) have observed a close associated between cpDNA fibrils and ribosome clusters in thin sections of the alga Ochromonas, though ribosome free nucleoid areas were observed. Though granules similar to the size of ribosomes are present in preparations of the current study they do not appear neatly spread along an RNA tanscript. It is possible that rather than this being a preparative difficulty that transcriptional complexes are RNP granules and some heterogenous RNP knobs (Fig.5) similar to some nonribosomal RNA transcripts observed in animal cell nuclei (e.g. Busby and Bakken, 1979; Hughes et aI., 1979; Puvion-Dutilleul and May, 1978). However, unlike chloroplasts, there is evidence in nuclei for RNP structures that are carriers of rapidly labelled extranucleolar RNA (Nash et aI., 1975). A further possibility to account for the granules in transcriptional strucures is that they represent RNA fortuitously associated with protein during isolation (Greenberg, 1975). However, it seems less likely that this would occur in the vesicle preparations. It should be noted that active gene visualization in membranous organelles has proven more difficult than with bacteria (Miller and Hamkalo, 1972; Laird et aI., 1973). On balance, the morphological studies suggest that in vivo, at least some nascent mRNA may be involved in translation, but more definitive approaches are required. In terms of chloroplast structure and organisation, our morphological observations suggest that there is thylakoid-associated chloroplast DNA extending into the stromal matrix in spinach chloroplasts, and it is associated with transcriptional complexes. The transcriptional complexes besides occurring in association with the thylakoids, would also contribute to the morphology of the stromal matrix. The purely DNA fibrillar areas (Ris and Plaut, 1962) may well be transcriptionally inactive. Acknowledgements The technical assistance ofJoan Brien and Margaret Gibberd is gratefully acknowledged. We wish also to thank the University of Newcastle Electron Microscope Unit. The project was funded by the Australian Research Grants Committee. Z. Pjlanzenphysiol. Bd. 106. S. 129-137. 1982.

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