To investigate the association between chloroplast DNA (cp DNA) and the photosynthetic membranes of spinach chloroplasts, previously suggested by electron-microscope autoradiography, use has been made of vesicles formed by isolating chloroplasts directly in 3·5 Mm Mg2+. These chloroplast vesicles consist of photosynthetic membranes, separate from chloroplast envelope membranes. Light and electron microscopy confirm that the vesicles consist of swollen stroma lamellar membranes with some peripheral grana lamellae that are much less swollen. Vesicles labelled with [3H]thymidine were obtained from [3H]thymidine-labelled chloroplasts from spinach disks in which chloroplast division and cp DNA synthesis and segregation were occurring. The chloroplast vesicle fraction retains about 45 % of the cp DNA as determined by liquid scintillation counting. The cp DNA-membrane associations do not appear to be dependent on the presence of Mg2+. The chloroplast vesicles can be autoradiographed for light microscopy if they are fixed in formaldehyde and no centrifugation steps are used. Lightmicroscope autoradiography is consistent with a preferential labelling of grana as opposed to stroma membranes, and long lengths of membrane are labelled. It appears that in spinach chloroplasts cp DNA is associated with granal thylakoids at intervals along the length of a continuous photosynthetic membrane system. Such an organization would facilitate cp DNA segregation during chloroplast division.

Chloroplasts contain DNA which codes for some of the proteins required for chloroplast development and function (Ellis, 1977; Kung, 1977). In higher plants there are many chloroplasts per cell, and chloroplast replication is associated with the full development of the leaf mesophyll cell (Possingham & Saurer, 1969). Some mechanism is required to ensure that chloroplast DNA (cp DNA) is transmitted to chloroplast progeny.

The chloroplast replication process can be conveniently studied in cultured spinach leaf disks, where, using autoradiography to detect [3H]thymidine incorporation into chloroplasts, it has been shown that chloroplast division is associated with cp DNA synthesis and an orderly transmission of the cp DNA to the daughter chloroplasts (Rose, Cran & Possingham, 1974; Possingham & Rose, 1976). From electronmicroscope autoradiography of sectioned chloroplasts, and chloroplast fractionation studies, it has been argued that it is the association of the cp DNA with the photosynthetic membranes that facilitates the cp DNA segregation process in spinach chloroplasts (Rose & Possingham, 1976a). Furthermore, rare light-microscope autoradiographs of whole chloroplasts showed continuous spiral labelling patterns which suggested that the cp DNA molecules occurred at intervals along the lamellar membrane system (Rose & Possingham, 1976 a). However, these latter autoradiographs could not be obtained routinely and a method using isolated photosynthetic membranes to autoradiograph the photosynthetic membrane system was sought. In this paper, chloroplast membrane vesicles have been used to study the association of cp DNA with the photosynthetic membrane system.

Incubation of cultured leaf disks with [3H]thymidine

Spinach (Spinacia olerecea L) leaf disks were obtained from the base of 2-cm-long leaves and cultured on sterile nutrient agar using the methods of Possingham & Smith (1972). The leaf disks were grown at 23 °C under fluorescent light of intensity 6 mW cm−2, with a 14-h day. Twenty to twenty five disks per plate were incubated in 50 μCi [6— 3H]thymidine (20-30 Ci/mM, Radiochemical Centre, Amersham, U.K.) added in 1 ml to the surface of the 20 ml of nutrient agar medium in 9-cm Petri dishes. All manipulations to the cultured disks were made in a sterile cabinet employing a laminar air-flow system. The incubation in [6— 3H] thymidine (3H—TdR) was continued for approximately 72 h (day o to day 3) which allows almost 3 cycles of chloroplast division, and ensures that all the cp DNA is extensively labelled (Possingham & Rose, 1976). In the experiment from which Figs. 16 and 17 were obtained, the incubation was from day 3 to day 6 where there would be about 2 cycles of division.

Isolation and cytology of chloroplasts and chloroplast vesicles

Intact chloroplasts were isolated in either 0·4 M sucrose + 3·5 mM MgCl2 (MgCl2 is subsequently referred to as Mg2+), or 0·35 M Mg2+, while chloroplast vesicles were isolated in 3·5 mM Mg2+. The disks were razor chopped in the appropriate medium to release the chloroplasts or vesicles, and then filtered through 70-μm steel mesh.

For light-microscope cytology the freshly isolated chloroplasts and chloroplast vesicles were examined with phase-contrast or Nomarski interference-contrast optics using a Leitz Orthoplan microscope.

For electron-microscope cytology a similar procedure to that of Fowke (1975) for isolated plant protoplasts was used. The isolated chloroplasts or vesicles were fixed in centrifuge tubes in their original isolation medium containing 1 % glutaraldehyde at pH 6·5 for 2 h at room temperature. Following the initial fixation, the chloroplasts or vesicles were pelleted at 1000 g and transferred to 3 % glutaraldehyde in isolation medium for a further 3 h. The isolated chloroplasts and vesicles were washed in isolation medium, then washed with 0·05 M sodium phosphate buffer at pH 6·8 at 0 °C. Postfixation in 1 % osmium tetroxide in 0·05 M sodium phosphate buffer at pH 6·8 at 0 °C was carried out overnight. The chloroplasts and vesicles were then washed with distilled water and dehydrated in an ethanol series at 0 °C, gradually transferred to pure propylene oxide in glass vials at room temperature, and finally embedded in Araldite in small alfoil trays. Thin sections were stained with uranyl acetate in 50 % ethanol and lead citrate before viewing in an AEI EM 6 G electron microscope.

Autoradiography

Light-microscope autoradiography was carried out with isolated chloroplasts and chloroplast vesicles following incubation of the spinach disks in [6— 3HJthymidine. Isolated chloroplasts were fixed in 4 % formaldehyde in 0·1 M sodium phosphate buffer at pH 7·2 at o °C for 20 min in test tubes, then transferred to 4% formaldehyde in 3·5 mM phosphate buffer at pH 7·2 for at least 12 h at 5 °C. Isolated chloroplast vesicles were fixed directly in 4% formaldehyde in 3’5 HIM phosphate buffer at pH 7·2 at o °C for 20 min, then transferred to 5 °C for at least 12 h. The chloroplasts or chloroplast vesicles sedimented by gravity to the bottom of the tube and the supernatant was removed and replaced with distilled water. The chloroplasts or chloroplast vesicles were pipetted on to a glass slide, air dried, and processed for autoradiography using Ilford L4 emulsion. The autoradiographic techniques have been described, as has the specificity of incorporation of 3H-TdR into DNA (Rose, Cran & Possingham, 1975). Following the development of the slides they were dehydrated in an ethanol series and mounted in Euparal.

Silver grains over the chloroplast vesicles were scored as to whether they were associated with grana-rich or stroma-rich membrane areas. Circular vesicles with a peripheral grana-rich region were scored. This did not exclude vesicles which had grana across the stroma membrane area or in the centre of the vesicle (Figs. 10-17). Vesicle diameter and stroma membrane diameters were measured. The stroma membrane area was calculated directly, and the grana membrane area by difference between the total vesicle area minus the central stroma area. Grains per unit area over the different membranes were obtained from too vesicles scored from slides from the same experiment as Figs. 10-15, but with an exposure period of 5-5 weeks. Background values were obtained adjacent to each vesicle scored, and corrections made for the counting of that particular vesicle.

[3H]thymidine incorporation into chloroplasts and chloroplast vesicles as determined by liquid scintillation counting

Chloroplasts or ‘chloroplast vesicles’ were isolated in the appropriate isolation medium at o °C as described earlier and immediately placed on a single step discontinuous sucrose gradient of 20 % and 60 % sucrose and centrifuged at 27 500 g with a sorvall HB-4 rotor for 25 min. The chloroplast fractions were collected from the 20 %/6o % interface and the sucrose diluted with distilled water to resuspend the fraction and allow it to be placed at the top of the 20 % sucrose. The fractions were then centrifuged and collected again using the same procedures as for the first centrifugation. With this procedure nuclei go to the bottom of the tube and are separated from the chloroplast fractions; it was routinely monitored with the light microscope.

After the double centrifugation procedure the chloroplasts and chloroplast vesicles were used to determine chlorophyll concentrations and the incorporation of 3H-TdR into trichloroacetic acid (TCA)-precipitable material.

Chlorophyll was determined by collecting an aliquot from the chloroplast fractions on a Whatman GF/A glass fibre filter. The filter was washed thoroughly with 3·5 mM Mg2+, then the chlorophyll extracted with 80 % acetone and estimated spectrophotometrically (Arnon, 1949).

The incorporation of 3H-TdR into TCA-precipitable material was determined by precipitation with 5 % TCA containing 0·5 mg/ml unlabelled thymidine. The precipitate was washed initially by centrifugation then collected on Whatman GF/A filters. The filters were washed with 5 % TCA, ether-ethanol, and finally ether before air drying and counting in a toluene-based scintillation fluid.

Dialysis experiments

Chloroplast vesicles free of nuclei were collected as described above and 1 ml of known chlorophyl concentration placed in dialysis tubing for dialysis against 1 1. of distilled water, or in the case of controls for dialysis against 3·5 mM Mg2+. Dialysis was carried out for 24 h. The dialysis solutions were changed twice. After dialysis the vesicles were pelleted at 1000 g and the 3H-TdR present in TCA-precipitable material was determined, in both treatments.

When chloroplasts from cultured spinach disks are isolated in 3·5 mM Mg2+ they form many structures similar to those in Figs. 1-3. It is well known that chloroplasts isolated in such hypotonic solutions lose their outer envelopes (Spencer & Wildman, 1962; Douce, Holtz & Benson, 1973). The structures formed in Figs. 1-3 are large vesicles formed from the photosynthetic membranes. I have referred to these structures as chloroplast vesicles.

Fig. 1.

Chloroplast vesicles isolated in 3·5 mM Mg2+. phase-contrast, × 1450.

Fig. 1.

Chloroplast vesicles isolated in 3·5 mM Mg2+. phase-contrast, × 1450.

The chloroplast vesicle structures are similar to those described by Spencer & Wildman (1962) when using 20 mM sucrose and lower concentrations as osmoticum. Subsequently, Spencer & Unt (1965) noted the ability of low concentrations of Mg2+ to form balloon-like structures and to stabilize grana and intergrana membranes relative to the disruption that occurs in distilled water. Magnesium was preferred for the study reported here as more and larger vesicles formed thanfrom a sucrose solution of equal osmotic pressure and they preserved more effectively for subsequent autoradiography. The interpretation of Spencer & Wildman (1962) based on lightmicroscope cytology is that the vesicles arise by swelling of the stromal membranes, and the grana which are more resistant to swelling, occur at the surface of these stromal balloons. This latter interpretation is consistent with the cytological observations of the current study. In Fig. 2 the grana appear as dark structures at the vesicle periphery, while using Nomarski interference-contrast optics the grana appear as small raised knobs (Fig. 3). Control chloroplasts isolated in 0·4 M sucrose +3·5 mM Mg2+ are shown in Figs. 4 and 5. In the chloroplasts of Fig. 4 no grana can be visualized, while they are just apparent in Fig. 5. Grana are visible in type II but not type I chloroplasts (Spencer & Unt, 1965). Type I chloroplasts are morphologically intact while type II chloroplasts have envelope breakage (Coombs & Greenwood, 1976) and were in the minority (about 20%) in control preparations from which Figs. 4 and 5 were obtained.

Fig. 2.

As for Fig. 1. × 3450.

Fig. 2.

As for Fig. 1. × 3450.

Fig. 3.

Chloroplast vesicle isolated in 3·5 mM Mg2+. Nomarski interference-contrast, × 3650.

Fig. 3.

Chloroplast vesicle isolated in 3·5 mM Mg2+. Nomarski interference-contrast, × 3650.

Fig. 4.

Chloroplasts isolated in 3·5 mM Mg2+ + o·4M sucrose. Nomarski interference-contrast. × 2800.

Fig. 4.

Chloroplasts isolated in 3·5 mM Mg2+ + o·4M sucrose. Nomarski interference-contrast. × 2800.

Fig. 5.

Chloroplasts isolated in 3·5 mM Mg2+ + o·4M sucrose. Nomarski interference-contrast. × 2800.

Fig. 5.

Chloroplasts isolated in 3·5 mM Mg2+ + o·4M sucrose. Nomarski interference-contrast. × 2800.

When the chloroplast vesicles are fixed in glutaraldehyde, processed, and examined in the electron microscope they appear as in Figs. 6-8. They are bound by a membrane to which adhere varying numbers of grana lamellae. The number of grana lamellae seen in a single section will vary according to the section plane. The grana are partially disrupted due to the isolation in the 3·5 mM Mg2+ and possibly due in part to the difficulty in preserving the vesicles for electron microscopy. Nevertheless, grana clearly are present at the vesicle periphery, and the vesicles form from stromal membranes. It is clear from electron microscopy and light-microscope observations of the vesicles that they are not uniformly covered with grana but there are a few granal rich areas adhering to the surface of the vesicles. An isolated intact chloroplast is shown in Fig. 9.

Fig. 6.

Electron micrographs of chloroplast vesicles isolated in 3·5 mM Mg2+. Fig. 6, × 11800;Fig. 7, × 1135o;Fig. 8, × 15500.

Fig. 6.

Electron micrographs of chloroplast vesicles isolated in 3·5 mM Mg2+. Fig. 6, × 11800;Fig. 7, × 1135o;Fig. 8, × 15500.

Fig. 7.

Electron micrographs of chloroplast vesicles isolated in 3·5 mM Mg2+. Fig. 6, × 11800;Fig. 7, × 1135o;Fig. 8, × 15500.

Fig. 7.

Electron micrographs of chloroplast vesicles isolated in 3·5 mM Mg2+. Fig. 6, × 11800;Fig. 7, × 1135o;Fig. 8, × 15500.

Fig. 8.

Electron micrographs of chloroplast vesicles isolated in 3·5 mM Mg2+. Fig. 6, × 11800;Fig. 7, × 1135o;Fig. 8, × 15500.

Fig. 8.

Electron micrographs of chloroplast vesicles isolated in 3·5 mM Mg2+. Fig. 6, × 11800;Fig. 7, × 1135o;Fig. 8, × 15500.

Fig. 9.

Electron micrograph of chloroplast isolated in 3·5 mM Mg2+ + o·4 M sucrose, × 26000.

Fig. 9.

Electron micrograph of chloroplast isolated in 3·5 mM Mg2+ + o·4 M sucrose, × 26000.

Fig. 10.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 10.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 11.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 11.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

The chloroplast vesicles can be fixed and processed gently with minimal manipulation for light-microscope autoradiography. When the vesicles dry down on to the slide only the grana rich regions can be readily visualized and the stroma appears clear. Observations indicate that the grana rich regions adhere first to the slide and then the remainder of the vesicle dries down. Many of these structures appear circular surrounded by grana with central stroma regions (Figs. 15 and 16).

Autoradiographs of chloroplast vesicles are shown in Figs. 10-15, and show grains following the vesicle periphery where there are granal rich regions. In Fig. 12 a line of grains extends towards the centre of the dried-down vesicle, it is likely that this is also a row of grana as at least some patterns like this will be seen, depending on how many granal areas there are and their distribution around the vesicle. Uniform labelling of the vesicles would show grains right across the vesicle, and would indicate stroma membrane labelling. Figs. 10-15 should be compared with Figs. 1-3.

Fig. 12.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 12.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Further autoradiographs of chloroplast photosynthetic membranes are shown in Figs. 16 and 17 from a separate experiment. The underlying membranes can be seen more readily in these photographs. Fig. 16 is the type of pattern shown in Figs. 1015 where there are peripheral grana rich regions. Fig. 17 shows a partially swollen chloroplast in which a number of swollen stromal regions are apparent, and the large vesicles free of central grana have not formed from this structure. The silver grains follow the relatively unswollen grana-rich regions, and not the swollen stromal regions. Most vesicles observed show the reported labelling patterns. Such patterns are not observed in the presence of the detergent Triton X-100 which solubilizes the chloroplast membranes. If vesicles are located on the slide then checked for incorporation, 84% are clearly labelled at the level shown in Figs. 10-17. The remaining 16% are virtually unlabelled.

To obtain a quantitative estimate of silver grain distribution in relation to grana and stroma membrane areas, circular vesicles with a peripheral grana area were scored as indicated in the Materials and methods section.

The grana membrane area is enriched in grana, but is not uniformly covered with grana (Figs. 6, 7, 11, 15). The stroma membrane areas are enriched in stroma membranes but may contain grana (Figs. 12, 17). The peripheral grana pattern could arise from a stroma membrane sheet interconnecting a row of grana on opposite sides of the sheet. On swelling a vesicle with an encircling ring of grana could be formed (e.g. Figs. 8, 9).

There are on average more than twice as many silver grains per unit grana-rich membrane area, compared to per unit stroma-rich area, and two thirds of the vesicle area is grana rich, giving four times as many grana grains (Table 1). A better indication of a preferential grana membrane labelling pattern, can be seen from the frequency distribution histograms of grains per unit area (Fig. 1). Thirty nine percent of the vesicles show less than 0·100 grainsm2 over stroma membranes compared with 1 % of grana membranes having this labelling density. It is these vesicles with the low stroma membrane labelling that have the clearest membrane separation pattern (Figs. 10, 11, 13-16), while the more heavily labelled stroma areas would likely contain grana (Figs. 12, 17). The range of values over the 2 membrane areas is fairly similar, it is the distribution which is quite different.

Table 1.

Silver grain distribution over chloroplast vesicles

Silver grain distribution over chloroplast vesicles
Silver grain distribution over chloroplast vesicles
Fig. 13.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 13.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

While the grana membrane areas have more membrane material because of their stacking, cp DNA would not be expected to occur between grana thylakoids. Electron microscopy (Rose & Possingham, 1976a), suggests that the cp DNA is membrane attached, and extends into the stroma matrix. It seems more likely that the labelled cp DNA is associated with the grana surface.

Autoradiographs of whole chloroplasts are shown in Figs. 18 and 19. These latter chloroplasts were isolated in 0·35 M Mg2+ and similar results were obtained from chloroplasts isolated in 0·4 M sucrose. When chloroplasts or chloroplast vesicles were dried-down on to slides for autoradiography, nuclei were also present in the preparation. Figs. 20 and 21 show that nuclei appear similar in autoradiographs of the 2 different types of treatment. Nuclei are much more heavily labelled than chloroplasts because of their much greater DNA content and are known to synthesize DNA in spinach leaf disks (Rose et al. 1975). The unlabelled circular region is the nucleolar region. Labelled nuclei can be clearly distinguished from chloroplasts.

Fig. 14.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 14.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 15.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 15.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Exposed 10 weeks, × 1900.

Fig. 16.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Separate experiment from that of Figs. 10-15. Exposed 6 weeks, × 3800.

Fig. 16.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Separate experiment from that of Figs. 10-15. Exposed 6 weeks, × 3800.

Fig. 17.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Separate experiment from that of Figs. 10-15. Exposed 6 weeks, × 3800.

Fig. 17.

Autoradiographs of 3H-TdR-labelled chloroplast vesicles isolated in 3·5 mM Mg2+. Separate experiment from that of Figs. 10-15. Exposed 6 weeks, × 3800.

Fig. 18.

Autoradiographs of 3H-TdR-labelled chloroplasts isolated in 0·35 M Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 18.

Autoradiographs of 3H-TdR-labelled chloroplasts isolated in 0·35 M Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 19.

Autoradiographs of 3H-TdR-labelled chloroplasts isolated in 0·35 M Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 19.

Autoradiographs of 3H-TdR-labelled chloroplasts isolated in 0·35 M Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 20.

Autoradiograph of 3H-TdR-labelled nucleus, isolated in 3·5 mM Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 20.

Autoradiograph of 3H-TdR-labelled nucleus, isolated in 3·5 mM Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 21.

Autoradiograph of 3H-TdR-labelled nucleus, isolated in 0·35 M Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 21.

Autoradiograph of 3H-TdR-labelled nucleus, isolated in 0·35 M Mg2+. Same experiment as Figs. 10-15. Exposed 10 weeks, × 1900.

Fig. 22.

Frequency distribution of silver grains/μm2 for stroma-rich (A) and grana-rich (B) membrane areas of chloroplast vesicles. Vesicles were isolated from tissue incubated 72 h (day o to day 3) in 3H-TdR.

Fig. 22.

Frequency distribution of silver grains/μm2 for stroma-rich (A) and grana-rich (B) membrane areas of chloroplast vesicles. Vesicles were isolated from tissue incubated 72 h (day o to day 3) in 3H-TdR.

In order to gain some indication of how much cp DNA remained associated with the autoradiographed chloroplast vesicles, the incorporated radioactivity in control chloroplasts and chloroplast vesicles was determined by liquid scintillation counting. Results are shown in Table 2. A variable but substantial amount of radioactivity was retained by the vesicles. Considering the shearing of cp DNA that would occur in vesicle formation the radioactive cp DNA must be tightly bound to the membrane. Clearly, however, it is fragments and not whole molecules that are being autoradiographed.

Table 2.

[3H]thymidine incorporation into chloroplasts and chloroplast vesicles

[3H]thymidine incorporation into chloroplasts and chloroplast vesicles
[3H]thymidine incorporation into chloroplasts and chloroplast vesicles

The question arises that the autoradiographs could represent indiscriminate cation binding of cp DNA. However the specific types of patterns obtained argue against this. Nevertheless, the possibility of indiscriminate binding was investigated by separating chloroplast vesicles from nuclei, and dialysing them against distilled water. Results are shown in Table 3. Following dialysis two thirds of the radioactive cp DNA still remained with the membrane fraction that could be pelleted at 1000 g. The loss of radioactivity that did occur is most likely due to further fragmentation of the membrane system that occurs in distilled water (Spencer & Unt, 1965), which would reduce membranes sedimenting at 1000 g. Indiscriminate Mg2+ binding of cp DNA to the membrane cannot account for the binding of most cp DNA to the chloroplast vesicles.

Table 3.

Dialysis of chloroplast vesicle fraction

Dialysis of chloroplast vesicle fraction
Dialysis of chloroplast vesicle fraction

By using chloroplast vesicles free of outer-envelope membranes, the photosynthetic membrane system of spinach chloroplasts can be autoradiographed directly, after having incorporated 3H-TdR during the growth of cultured spinach disks. From the data obtained it can be argued that what are being autoradiographed are fragments of cp DNA attached to the photosynthetic membranes. When the chloroplast envelope breaks, and is lost, after chloroplasts are isolated in 3·5 mM Mg2+, substantial cp DNA is retained by the thylakoid membranes. The 45% label retention obtained in this study, compares with a 61 % value obtained (Rose & Possingham, 1976a) using isolated chloroplasts swollen in 4 mM Mg Cl2 but also containing 10 mM buffer, which causes less swelling of thylakoid membranes (Douce et al. 1973). Furthermore, cp DNA and membrane associations have been observed by electron microscopy in osmotically shocked spinach chloroplasts (Woodcock & Fernández-Morán, 1968).

The chloroplast vesicle cytology and autoradiography provides evidence that is consistent with 2 points. First, that cp DNA occurs along the photosynthetic membrane system (e.g. in Figs. 14 and 17 about 29 and 36 μm respectively of membrane are labelled) and secondly, that cp DNA is preferentially associated with the grana lamellae (Table 1, Fig. 1).

The significance of these results for spinach chloroplasts lies in providing an explanation for the segregation of cp DNA in an equitable way in the dividing chloroplast. That dividing spinach chloroplasts do segregate their DNA in a fairly equal way to daughter progeny has been shown using pulse-chase experiments in cultured spinach disks (Rose et al. 1974).

The electron-microscope autoradiography data obtained with thin sections (Rose & Possingham, 1976a) and the current study using light-microscope autoradiography of whole, large vesicles is most readily explained by the cp DNA occurring at intervals along a continuous photosynthetic membrane system, attached to the grana lamellae. That the photosynthetic membrane system is a continuum is the current understanding of the chloroplast morphology of higher plants (Coombs & Greenwood, 1976; Gunning & Steer, 1975). There are some recent indications from observations by light microscopy of living chloroplasts (Jope, Atchison, Pringle & Wildman, 1977) and from light microscopy using a fluorescent dye staining for DNA (Jope, James & Wildman, 1977) that are also consistent with cp DNA occurring along the photosynthetic lamellar system of chloroplasts.

A preferential association of cp DNA with the grana, raises the question of how many granal stacks have an associated cp DNA molecule. Overall the chloroplast vesicle studies suggest that all grana could have associated cp DNA as most vesicles are extensively labelled even though some label is lost during preparation. However, unlabelled grana regions do occur, and 16% of the vesicles are unlabelled. It is of some value to consider that higher plant chloroplasts usually have 20-30 cp DNA copies (Kirk, 1972) and 40-60 grana (Gunning & Steer, 1975).

It should be emphasized that in the spinach disk system studied, there is a regular growth and division cycle, coupled to cp DNA synthesis and segregation (Rose et al. 1974; Possingham & Rose, 1976). It is therefore possible that cp DNA replication is coupled to membrane growth in a way similar to that postulated for bacteria i.e. that chromosome duplication and separation is coupled to an extension of the membrane surface (Ryter, 1968). Evidence has also recently been presented for the association of mitochondrial DNA at or near its origin of replication, with the inner mitochondrial membrane (Albring, Griffith & Attardi, 1977). A role in mitochondrial DNA replication and or segregation is suggested. Whether in higher plant chloroplasts there is additional significance for cp DNA being associated with the granal membranes (e.g. in granal development), rather than other membranes such as the inner envelope is unclear. Bisalputra & Bisalputra (1969) have suggested that the contact between the nucleoid and photosynthetic lamellae in the brown alga Sphacelaria may be significant in the growth of the chloroplast membrane system.

Herrmann (1970) and Kowallik & Herrmann (1972) have studied the organization of cp DNA in the chloroplasts of higher plants using Beta vulgaris L. They have concluded that cp DNA in mature chloroplasts is distributed within several nucleoids (depending on chloroplast size) with each nucleoid containing 4-8 genetic units. The degree of clustering of cp DNA molecules might depend on the degree of coupling between cp DNA replication and membrane growth, which would be influenced by chloroplast development and physiology. In this regard it should be noted that cp DNA synthesis and plastid division are readily separated experimentally i.e. cp DNA synthesis without plastid division (Rose et al. 1975; Kass & Paolillo, 1977) and plastid division without cp DNA synthesis (Boasson & Laetsch, 1969; Rose & Possingham, 1976b).

The association of cp DNA with thylakoids is also important in cp DNA segregation in the brown alga, Sphacelaria (Bisalputra & Bisalputra, 1970). Sphacelaria has a ring nucleoid (Bisalputra & Bisalputra, 1969) attached to the inner thylakoids of the peripheral lamellae which loop around the rim of the chloroplast (Bisalputra & Bisalputra, 1969; Bisalputra & Burton, 1970). A similar situation exists in Ochromonas (Gibbs, Cheng & Slenkis, 1974) where it is known that there are a number of cp DNA molecules, which are probably distributed along the nucleoid (Gibbs & Poole, 1973; Gibbs et al. 1974).

Chloroplasts of higher plants (Rose & Possingham, 1976a) and many algae (see literature cited by Gibbs et al. 1974) when studied by electron microscopy have nucleoids apparently scattered throughout the chloroplast. It is possible that in such chloroplasts the situation is similar to that in spinach, where there appears to be a specific organization of cp DNA in relation to the photosynthetic membrane system.

I wish to thank Margaret Gibberd for her valuable assistance throughout the study, and Margaret Brosnan for assistance with sectioning for electron microscopy. The project was supported by ARGC grant No. D2-76/15280.

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