Richard A. White and J. Kenneth Hoober'. Department of Botany and ... tional state of PSI[, changed only slightly during the first 2 h of greening. When these cells ...
Plant Physiol. (1994) 106: 583-590
Biogenesis of Thylakoid Membranes in Chlamydomonas reinhardtii y l
'
A Kinetic Study of Initial Greening Richard A. White and J. Kenneth Hoober'
Department of Botany and Center for the Study of Early Events in Photosynthesis, Arizona State University, Tempe, Arizona 85287-1601
Chl a and b accumulate without a lag when dark-grown cells of Chlamydomonas reinhardtii y l are exposed to light at 38OC (Maloney et al., 1989). Accumulation of the Chl a/bbinding proteins (LHCP) of LHCII parallels the increase in Chls, and Chl b is quantitatively incorporated into newly assembled LHCs (Hoober et al., 1990). An increase in PSII activity, which is negligible in dark-grown cells, also is initiated by light (Hoober et al., 1991). Micrographs of rapidly greening cells show that these biochemical events are accompanied by development of membrane structures that appear to flow from localized regions of the chloroplast envelope (Hoober et al., 1991).The kinetics of these processes, coupled
with the rapid degradation of LHCP in the dark (Hoober et al., 1990), presumably by a membrane-bound protease (Hoober and Hughes, 1992), suggests that biogenesis of thylakoid membranes in this organism is initiated at the level of the chloroplast envelope by light-activated Chl synthesis. Evidence that envelope membranes are a major site for synthesis of thylakoid lipids also supports this suggestion (see Hoober et al., 1994, for review). Our purpose in developing the greening of C. reinhardtii y l as an experimental system was to take advantage of the initial kinetics of Chl synthesis and thylakoid biogenesis when the cells are exposed to light at slightly elevated temperatures (Hoober and Stegeman, 1976; Maloney et al., 1989). The higher temperature induces expression of some genes, the lhcb genes in particular, in the dark and ensures a supply of LHCP upon illumination. Similar effects of heat on expression of light-inducible genes have been observed in barley and pea seedlings (Otto et al., 1992; Beator et al., 1993) and may be a general phenomenon. v. Gromoff et al. (1989) observed that heat-shock genes in Chlamydomonas were inducible by light, which is the corollary of light-inducible genes induced by elevated temperature (Hoober et al., 1982; Kindle, 1987). Incubation of C. reinhardtii cells at 38OC is not detrimental to protein synthesis or chloroplast development (Hoober and Stegeman, 1976; Hoober et al., 1991). After incubation in the dark at 38OC, exposure of cells to light initiates assembly of photosynthetic units by providing the remaining essential component, Chl. Whether synthesis of Chl or Chl-binding proteins is rate limiting under these conditions is not known. This system may permit these questions to be approached, and assembly of the complexes investigated, in real time. An analysis of initial kinetics of assembly of protein-Chl complexes may also be useful in distinguishing between assembly at the level of the envelope or at sites deeper within
' This work was supported by grant DCB-0018797 from the National Science Foundation. This is publication 198 of the Arizona State University Center for the Study of Early Events in Photosynthesis. The Center is funded by Department of Energy grant DEFG02-MER13969 as part of the U S . Department of Agriculture/ Department of Energy/National Science Foundation Plant Science Centers program. * Correspondingauthor; fax 1-602-965-6899.
mal fluorescence; Fo, fluorescence under conditions of minimal reduction of QA;Fp, peak fluorescence; Fs, steady-state fluorescence; FT, fluorescence 300 ms after start of actinic light; Fv, variable fluorescence (FM - Fo); LHCI (LHCII), light-harvesting complexes of PSI (PSII); LHCP, light-harvesting Chl a/b-binding apoprotein of LHCII; QA(Qs), plastoquinonein the binding sites of the PSII reaction center apoproteins D2 (Dl).
Initiation of thylakoid membrane assembly was examined in degreened cells of Chlamydomonas reinhardfii yl cells depleted of thylakoid membranes and photosynthetic activity by growth in the dark for 3 to 4 d. Photoreductive activities of photosystem II (PSII) and photosystem I (PSI) increased with no apparent lag when degreened cells were exposed to light at 38'C. However, fluorescence transients induced by actinic light, which reflect the functional state of PSI[, changed only slightly during the first 2 h of greening. When these cells were treated with 3-(3,4-dichlorophenyl)-1,l-dimethyl urea (DCMU) or saturating light, fluorescence increased commensurate with the cellular content of chlorophyll. In similar experiments with greening cells of C. reinhardfii CC-2341 (ac-u-g-2.3), a PSI-minus strain, fluorescence increased with chlorophyll without treatment with DCMU. These data suggested that fluorescence of initial PSI1 centers in greening y l cells was quenched by activity of PSI. Continuous monitoring of fluorescence in the presence or absence of DCMU showed that assembly of quenched PSI1 centers occurred within seconds after exposure of yl cells to light. These results are consistent with initial assembly of PSI and PSI1 within localized domains, where their proximity allows efficient energy coupling.
Abbreviations: DCPIP, 2,6-dichlorophenolindophenol;FM, maxi-
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the organelle. One of several kinetic processes may limit assembly of LHCII: (a) The rate of transport of the LHCP precursor across the envelope may be limiting. (b) The rate of integration of LHCP into thylakoid membranes and/or association with Chl may be limiting. These two events may be indistinguishable if assembly occurs at the level of the envelope. (c) Integration of LHCP into thylakoids may occur at sites within the organelle. In this case, diffusion of LHCP through the stroma and buildup of a sufficient pool to sustain a high rate of integration may be required, which possibly can be detected as a unique kinetic pattem. From studies with isolated chloroplasts (Reed et al., 1990), the time for diffusion through the stroma is expected to be in the minute range. To investigate these possibilities, it is necessary to examine the initial kinetics of greening, i.e. the rate immediately after exposure of cells to light. As a continuation of our work on thylakoid membrane biogenesis, we used the fluorescence properties of PSII to assess the initial kinetics of thylakoid biogenesis in C. reinhardtii. The small amount of Chl in degreened cells is highly fluorescent (Hoober et al., 1991), which indicated that residual LHCIIs either are not connected to reaction centers (Cahen et al., 1976) or are associated with centers unable to transfer electrons from QA to Qe (Guenther et al., 1990; Neale and Melis, 1990). We also observed that the steady-state room temperature fluorescence of intact cells or membranes did not increase significantly as greening progressed (Hoober et al., 1991). This quenched state of newly synthesized Chl was the basis for the experimental approach. MATERIALS AND METHODS
Cells
Chlamydomonas reinhardtii yl used in these experiments was a line maintained in this laboratory. Strains FUD34 (CC2518) and F34 (CC-2340) were provided by Dr. Elizabeth H. Hams from the Chlamydomonas Genetics Center, Duke University. Strain CC-2341 (ac-u-g-2.3) was kindly provided by Dr. Andrew Webber (Department of Botany, Arizona State University). Except for F34, which did not grow in the dark, these strains were phenotypically y when grown in the dark. Greening experiments were performed as described previously (Maloney et al., 1989). Fluorescence Induction Kinetics
Greening cells were collected by centrifugation and resuspended in 50 mM Tricine-KOH (pH 7.4) containing 5 m MgC1’. Samples were dark adapted for at least 5 min before exposure to modulated measuring light of low intensity (2.5 pmol photons m-’ s-I) for determination of FO and then to actinic light (80 pmol photons m-’ s-I) to obtain fluorescent transients with the PAM Chl fluorescence monitor (Heinz Walz GmBH, Effeltrich, Germany). F M was determined with saturating pulses (0.5 s) of white light (1200 pmol photons m-’s-’) or by addition of DCMU to a concentration of 10 p ~ Chl . concentration in samples was adjusted to 10 &mL or less to avoid reabsorption of fluorescent light. This protocol gave reproducible Fo and quantum yield values for green
Plant Physiol. Vol. 106, 1994
cells that were comparable to those reported in the literature for algal cells (Biichel and Wilhelm, 1993). Continuous Monitoring of Fluorescence in Greening
of yl Cells Suspensions of cells preincubated in the dark at 38OC for 1.5 h were transferred to the sample cuvette of the PAM fluorescence monitor maintained at 38OC. After the sample was stirred for 2 min in the dark, fluorescence was monitored continuously, with measuring and actinic light applied concurrently (total intensity, 30 pmol photons m-’! s-’). DCMU was added to 10 PM either immediately before or during the illumination period. Following illumination, Chl concentration in 80% acetone extracts of cell suspensioiis was determined fluorometricallyor as described by Porra et al. (1989). Photosystem Assays
PSI1 activity was assayed by the change in Asg0 of the reaction mixture that resulted from photoreduclion of DCPIP (emM = 16 cm-’) as described previously (Hoober et al., 1991). PSI was assayed by photoreduction of methyl red (Vemon et al., 1966; Hoober et al., 1969).The reaction mixture contained 10 ~ lNa2HP04, l ~ 5 m Tricine-KOH, 5 m KCl, 0.5 m MgC12, 10 PM DCMU, 0.1 m DCPIP, 0.14 rmd methyl red, and cell extract routinely equivalent to lo7 cells/mL. N2 was bubbled through reaction mixtures for 2 min bcfore addition of ascorbic acid to 2.8 m (Hoober et al., 1969), which provided a final pH of 7.4. The stream of N2 was continued as cell extract was added and the incubation mixture was pumped through a flow cell to measure the change in A430 resulting from reduction of methyl red (emM = 1 2 cm-I). After a dark rate was established, the mixture was exposed to light from a tungsten-halide lamp that was passed through glass IR (Coming glass No. 4602) and 640-nm cutoff (Coming glass No. 2418) filters. The rate of photoreduction was linear with respect to amount of extract added. PSI and PSII were assayed with the same fluence of incident light (400 pmol photons m-’ s-I) that provided approximately 50% maximal velocity of PSII. 77 K Fluorescence
Greening cells were suspended in g1ycerol:growth medium (2:1, v/v) and frozen in liquid Nz. Emission spectra were recorded with a FluroMax spectrofluorometer (SPEX Industries, Inc., Edison, NJ) at an excitation wavelength of 438 or 472 nm. Slit widths were 1.0 and 0.25 mm for the excitation and emission monochromaters, respectively. RESULTS Fluorescence Characteristics of Greening Cells
Shown in Figure 1 are fluorescence induction transients typically obtained with fully greened cells, used as control samples. The fluorescence yield increased within 300 ms in actinic light to Fp and declined thereafter to an Fs level, which described a typical Kautsky effect. Addition of DCMU to the cell suspension, which blocks electron flow from QA to QB,
Kinetics of PSI1 Assembly in Chlamydomonas
was somewhat slower after 1 h of greening (compare curves for samples without DCMU in A and B of Fig. 2), the increase was approximately of the same magnitude in both samples. As illustrated in Figure 2B, fluorescence of the sample often decreased slightly at the time (300 ms) when Fp was reached in green cells (Fig. 1).As a result, Fo and FT values did not increase proportionally with the 4- to 8-fold increase in Chl
1.5 2
Fm
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>
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A
0.5L
I
2 1
0.4
I
0.4
0.75
Time
I
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Figure 1. Fluorescence induction kinetics displayed by green C. reinhardtii y l cells. Cells were suspended in 50 mM Tricine-KOH (pH 7.4) containing 5 mM MgClz to a density of 2 X lob celk/mL.
2
Fluorescence was measured at room temperature in the absence (curve 1 ) or presence (curve 2) of 10 p~ DCMU. FpI, The initial plateau level of the fluorescence transient.
caused a greater and more rapid increase to FM. This pattern is characteristicof mature plant cells and has been extensively studied in C. reinhardtii, other algae, and higher plants (Guenther et al., 1990; Neale and Melis, 1990; Krause and Weis, 1991; Falk et al., 1992; Büchel and Wilhelm, 1993). Exposure of degreened cells to actinic light caused a rapid but small increase in fluorescence within 100 ms (Fig. 2A). Addition of DCMU caused only a slight, if any, increase in fluorescence yield or rate at which FM was reached. To investigate further the source of this small Fv signal, we examined greening cells of strain FUD34, which accumulate LHCII but not stable PSII centers because of a mutation in the chloroplast psbC gene (Rochaix et al., 1989). No increase in fluorescence yield in response to actinic light was observed with FUD34 cells, although fluorescence increased along with the amount of Chl during greening (data not shown). Furthermore, green cells of the nuclear mutant F34, which is phenotypically similar to FUD34 (Rochaix et al., 1989; de Vitry et al., 1991), showed no Fv signal, as described previously (Wollman and Delepelaire, 1984; Girard-Bascou et al., 1992).The results with y l cells shown in Figure 2A suggested, therefore, that at least some of the residual LHCs were associated with reaction centers. However, the negligible PSII activity in degreened cells indicated that these centers lack the ability to reduce QB, which is characteristicof PSII centers in dark-grown Chlamydomonas cells (Cahen et al., 1976; Guenther et al., 1990). After the cells had greened for 1 h, the fluorescence induction transient was essentially unchanged from that at the beginning of greening. Although the increase in fluorescence
_-
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Figure 2. Fluorescence induction kinetics displayed by greening y l cells. Kinetics were determined as for Figure 1 except that the cell density was 5 X 10b/mL.A, Degreened cells before exposure to light; 6, greening cells exposed to light for 1 h; C , greening cells exposed to light for 1 h and then broken by passage through a French pressure cell at 5 K p.s.i. For each panel, fluorescence was measured with t h e same sample first in the absence (curve 1) and then in the presence (curve 2) of 10 p~ DCMU.
White and Hoober
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during 1 or 2 h of greening, respectively, and no Kautsky effect was apparent. (We refer to the fluorescence yield 300 ms after the start of actinic light as FT, i.e. fluorescence at the time at which Fp normally occurs [see Fig. 11.) Extending the time beyond that shown in Figure 2 did not reveal evidence of a further, delayed increase in fluorescence. However, FM and Fv/FM, which has been used as an estimate of the photochemical efficiency of PSII (Krause and Weis, 1991; Falk et al., 1992), increased with time of greening (Fig. 3). The increase in Fv/FM indicated that functional PSII centers were generated during this period, as expected from the increase in PSII activity (Hoober et al., 1991; see Fig. 4). With cells greened for 3 h or more, Fv/FM approached 0.7 (data not shown), near the value for green cells shown in Figure 1. The increase in F M but not F T during greening suggested that the initial PSII centers were unable to generate a more reduced state of QAwith relatively low-intensity actinic light (80 pmol photons m-’ SKI). In contrast, with membranes from greening cells, actinic light alone increased fluorescence yield to near F M (Fig. 2C). Dilution of electron acceptors such as NADP+ in broken-cell samples apparently reduced the rate of oxidation of the plastoquinone pool. The normal increase in fluorescence of membrane samples suggested, therefore, that quenching of PSII fluorescence in intact greening cells was not the result of assembly of atypical PSII centers or an excessively large pool of plastoquinone. Rather, efficient trapping of energy possibly was responsible for the quenching of the newly assembled units. If so, the capacity for electron transfer from PSII to PSI may have been established during the initial stage of greening. Quenching, then, should be eliminated in a PSIdeficient mutant strain. To evaluate this suggestion, we examined fluorescence of
Plant Physiol. Vol. 106, 1994
25 IC
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aal o
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Figure 4. Change in PSI and PSI1 activities during greening of yl
cells. Cells were suspended in 50 m M Tricine-KOH (pH 7.4) containing 5 m M MgCI2 and broken by passage through a French pressure cell at 5000 p.s.i. Aliquots were used to assay PSI (curve 1) by photoreduction of methyl red (MR) or PSI1 (curve 2) by photoreduction of DCPIP at the same incident light intensity. The results are mean values from three experiments.
mutant strain CC-2341, which contains a frameshift mutation in psaB, the chloroplastic gene encoding one of the subunits
of the PSI reaction center, and is devoid of PSI activity (Webber et al., 1993). Fluorescence of CC-2341 cells increased in parallel with Chl during 2 h of greening. h i contrast to results with yl cells, fluorescence of greening CC-2341 cells increased to F M with only actinic light (80 pmol photons m-’ s-l) similar to other PSI-deficient mutants (Webber et al., 1993). Addition of DCMU increased the rate at which FM was achieved but did not increase the fluorescence yield above the final steady-state level obtained with actinic light alone (data not shown). Appearance of PSI
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Figure 3. Fluorescence parameters determined with greening yl cells. Fo (O, curve 1) and FT (W, curve 2) were determined with greening cells, as shown in Figure 2, A and B, in the absence of DCMU. FyIFM (O, curve 3) and FM (O, curve 4) were determined
with saturating pulses of white light. Fo,FT, and F M are expressed in volts. In separate experiments, the patterns of the transients were the same, although the initial level of Chi varied between 1 .O and 1.5 pg/107 cells. The results are mean values from five experiments.
Biochemical assays showed that PSII activity increased approximately in parallel with Chl when degremed yl cells were exposed to light (Hoober et al., 1991). lis shown in Figure 4,activity of PSI also increased rapidly during the first 30 min of greening in yl cells but lagged behind the increase in PSII thereafter. After 3 h of greening, the ratio of activity of PSI to that of PSII was 0.4 to 0.5, similar to that found in green cells by these assays. The 77 K fluorescence spectrum of degreened yl cells contained a maximum at 678 nm but lacked the emission at 715 nni (Fig. 5) that is characteristic of LHCI in Chlamydomonas (Gershoni and Ohad, 1980; Delepelaire and Wollman, 1985).An increase in fluorescence at 715 nm was not detected until after 15 min in the light and then increased rapidly (Fig.
587
Kinetics of PSI1 Assembly in Chlamydomonas
--
Figure 5. Fs spectra at 77 K of greening yl cells. Spectra were determined after exposure of cells to light for O (curve I), 1 (curve 2), or 2 h (curve 3) at 38°C or 2 4 h (curve 4) at 25°C. Spectra were normalized to the maximal value in curve 1. Inset, Time course of maximal emission (curve 1) of yl cells at 715 n m or (curve 2) of CC-2341 cells at 708 nm.
41
u)
Time
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5 , inset). When the cells had been in the light for 2 h, the relative fluorescence of PSI was similar to that of green cells. Experiments with strain CC-2341 showed that fluorescence intensity at 708 nm, the emission maximum of LHCI at 77 K in the absence of PSI reaction centers (Webber et al., 1993), also increased after a lag at a rate similar to that of the 715nm fluorescence in y l cells (Fig. 5, inset). Initial Kinetics of PSI1 Assembly
The maximal 77 K fluorescence emission of PSII antenna at 678 nm in degreened cells was characteristic of free LHCII (Gershoni and Ohad, 1980; Krause and Weis, 1991; Hemelrijk et al., 1992). The peak emission shifted toward longer wavelengths with time in the light and was 683 nm after 1 h, 685 nm after 2 h, and 688 nm in fully green cells (Fig. 5 ) . This red shift possibly indicated progressive assembly of PSII units, with an increasing contribution from Chl a-protein complexes of the PSII core that in Chlamydomonas contain apoproteins of approximately 47 kD (Ae,,, = 685 nm) and/or 50 kD (Ae,,, = 695 nm) (Krause and Weis, 1991). The initial 678-nm peak did not persist, which suggested that residual LHCII in degreened cells were incorporated into new PSII units. Alternatively, the initial shift to 683 nm may indicate association of LHCII with PSI, a state-2 condition that yields peak emission at 682 nm (Wollman and Delepelaire, 1984). These data suggested that complete assembly of PSII units occurred relatively slowly during 1 to 2 h of greening. However, photoreductive activity increased without a lag when y l cells were exposed to light (Fig. 4). To determine the kinetics of initial greening, i.e. during the first few minutes in the light, it was necessary to overcome variations in sampling and low signal-to-noise ratios in measurements of biochemical activities or fluorescence induction within the first 5 min of greening. For this purpose, fluorescence of the
same cell suspension was monitored continuously at 38OC. The light-emitting diode in the PAM fluorescence monitor has a maximal output at 650 nm, the wavelength that supports optimal Chl synthesis in Chlamydomonas cells at 38OC (Hoober and Stegeman, 1976). Because Fs did not increase during the 1st h of greening of y l cells in white light (Fig. 3), we tested whether initial kinetics of assembly of PSII could be assayed by the difference in fluorescence in the absence or presence of DCMU, which does not impair initial greening (Hoober et al., 1991). As shown in Figure 6, fluorescence of cells greening under 650-nm light (30 pmol photons m-' s-') did not increase with time above the initial level until DCMU was added. When DCMU was added at the time of exposure to light, however, fluorescence intensity increased continuously. The 20 to 30% increase in fluorescence intensity during a 5-min period corresponded to the increase in Chl expected from the rate of accumulation described previously (Maloney et al., 1989) and confirmed by spectrofluorometric measurements of Chl in 80% acetone extracts of cells at the end of the illumination period (data not shown). The cells used in these experiments contained an initial Chl level of about 1 pg/107 cells and typically accumulated 3 to 4 pg Chl/107 cells during 1 h of greening (Maloney et al., 1989). As these experiments demonstrate, the rate of assembly of membranes can be studied kinetically during a period of a few minutes. DISCUSSION
Chl synthesis and thylakoid formation have been studied extensively in higher plants and green algae (see Hoober, 1987; Hoober et al., 1994, for reviews). Chloroplast development under temperate conditions requires expression of essential genes, particularly in the nucleus, that are regulated by light. The ensuing lag period of several hours after onset of illumination seems largely to reflect the time course of
White and Hoober
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1 electron flow through Cyt
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Continuous monitoring of fluorescence in greening yl cells. Suspensions of degreened cells were incubated at a density of 5 x IO6 cells/mL in the dark at 38°C for 1.5 h and then transferred into the measuring cuvette of the PAM Chl fluorescence monitor, which was operated in the continuous monitoring mode. After an additional 2 min in the dark, the measuring and actinic light were turned on simultaneously. Curve 1 shows fluorescence in a sample to which DCMU was added to 10 ~ L L Mafter 3 min in the light. For curve 2, DCMU was added immediately before cells were exposed to light. The traces show the extent of noise relative to the change in fluorescence at this early time of greening. Figure 6.
induction of these genes (Hoober et al., 1982; Malnoe et al., 1988; Anandan et al., 1993). The rate-limiting process during the initial phase of membrane formation, therefore, is likely synthesis of the gene products, many of which bind Chl. Subsequent steps, such as insertion of the proteins into membrane and association with Chl, probably require seconds or minutes and thus are more rapid than the rate of induction of gene expression. In the experiments described in this paper, the unique fluorescence properties of thylakoid membrane complexes were used to examine the initial kinetics of assembly under conditions in which induction of gene expression is not the limiting factor. The data obtained in this study show that fluorescence of PSII units assembled during the initial minutes of greening is strongly quenched. Several mechanisms may be responsible for this phenomenon: (a) Chlamydomonas cells exhibit a significant rate of chlororespiration (Peltier and Schmidt, 1991), which may quench fluorescence of the initially small number of PSII centers by maintaining the plastoquinone pool in an oxidized state. In Chlamydomonas thylakoids, as in photosynthetic bacteria and cyanobacterial membranes, respiratory and photosynthetic electron transport pathways share intermediates, in particular the quinone pool (Scherer, 1990). In isolated membrane fractions, effects of an active chlororespiration pathway that consumes reducing equivalents generated by PSII should still be apparent. However, PSII centers were not quenched in membrane fractions from y l cells. Although chlororespiration may be involved in quenching fluorescence, it is likely not the major factor. (b) Cyclic
Plant Physiol. V8A.106, 1994
b559 (Buser et al., 1902; Nedbal et al., 1992) possibly traps energy within PSII centers. This possibility seems unlikely because no quencb ing was observed in greening CC-2341 cells that have PSI[ but lack PSI activity. (c) High light intensities, in excess of that which is efficiently utilized by PSII units, promote de-e poxidation of violaxanthin to zeaxanthin (Demmig-Adams and Adams, 1992). In conjunction with an energy gradient across thylakoid membranes, zeaxanthin may cause thermal dissipation of absorbed energy and thus quench PSII fluoi.escence nonphotochemically (Gilmore and Yamamoto, 1993; Ruban et al., 19'94).Under ambient 0 2 and C 0 2 concentrations, maximal de-epoxidation in higher plants occurs within 10 to 15 min at (white) light fluences greater than 1000 umol photons m-2 s-l (Demmig-Adams et al., 1990; Jahns and Krause, 1994). Recovery of photosynthetic activity in spinach occurs at a fluence of 52 pmol photons m-2 s-' (Demmig-Adams and Adams, 1993). The relatively low fluence of (red) light of 30 pmol photons m-' s-' that supported greening of C. reinhardtii yl cells should not cause significant de-epoxidation of violaxanthin. Thus, nonphotochemical quenching or reduction in fluorescence yield as the result of photoinhibition (Krause and Weis, 1991) seems unlikely to account for the essentially complete quenching of fluorescence during the first few minutes of greening, although subsequent formation of small vesicles (Hoober et al., 1991)could lead to conditions suitable to maintenance of a pH gradient. (d) Quenching of fluorescence of PSII may result from assembly of PSI and PSII within localized sites that allow close interaction between the two centers. Trissl and Wilhelm (1993) argued that when PSI is near PSII, i.e. when the centers arc? energetically coupled, the kinetically more rapid PSI center!; drain excitation energy from PSII, thereby quenching its fluorescence. Within the early thylakoid membranes, LHCII may transfer energy to both PSII and PSI, a state-2 condition. The role of stacking as thylakoid membranes increase in ainount may be to separate the two photosystems and enhance the efficiency of PSII (Trissl and Wilhelm, 1993). Studies Jf chloroplast development have shown that stacking doe,c not seem to occur until an extensive amount of thylakoids has been made (Ohad et al., 1967b; Robertson and Laetsch, 15174;Hoober et al., 1991). In the first minutes of greening of C. reinhardtii y l , therefore, quenching of PSII fluorescence may result primarily from these centers being coupled to PSI. Although fluorescence spectra and biochemical assays indicated a low activity of PS1 in degreened cells, these assays perhaps underestimate this activity. Our results at least suggest that PSI activity is sufficient to drain electrons from the plastcbquinone pool faster than PSII can reduce it. Electron camcrs such as Fd and Cyt f remain at high levels in degreened jl cells (Ohad et al., 1967a) and may facilitate electron flow through PSI. The relatively high PSI activity required for quenching of PSII fluorescence in greening C. reinhardtii, yet with an apparent brief lag in accumulationof LHCI (Fig. 5), suggested that energy was initially transferred into PSI centers from LHCII. An analogous situation exists in Prochlorothrix hollandica, in which the photosystems normally are lot segregated (Post et al., 1993). Light of 650 nm, such as that used in our analysis of fluorescence induction kinetics, promoted a state-
Kinetics of PSI1 Assembly in Chlamydomonas
1 to state-2 transition during a period of 15 to 20 min in Prochlorothrix, and the resulting fluorescence induction transient was similar to that obtained with greening Chlamydomonas (Fig. 2B). We conclude that the initial organization of PSII and PSI in Chlamydomonas thylakoid membranes is possibly similar to that which occurs in Prochlorothrix exposed to red light. The kinetics of increase in fluorescence in the presence of DCMU (Fig. 6) suggested that complete PSII units were assembled within seconds after exposure of cells to light. Although in the first few minutes of greening the total number of PSII centers is low, they are apparently in sufficiently close proximity to PSI centers for efficient exciton or electron transfer. This conclusion assumes that the initial increase in PSII activity depends on association with LHCII, which is the predominant repository of newly synthesized Chl (Hoober et al., 1991). The absence of a detectable lag in assembly of PSII units is not consistent with the requirement for accumulation of a pool of LHCP in the stroma, with subsequent targeting and integration into membrane structures within the chloroplast. Rather, these results support the possibility that local domains of thylakoid membrane are assembled within the structures emanating from chloroplast envelope of cells greened for only a few minutes as revealed by EM (Hoober et al., 1991). A flow of membrane as vesicles from the envelope to the chloroplast interior may carry lipids and proteins originally on the envelope to developing thylakoids, if this pathway is a general feature of initial chloroplast development as proposed previously (Hoober, 1987; Hoober et al., 1994). Secondary sites of membrane assembly may thereby result as chloroplast development proceeds. This possibility is supported by insertion of LHCP into thylakoid membranes purified from partially developed pea a n d barley chloroplasts when incubated with stromal proteins and ATP (Cline, 1986; Chitnis et al., 1987). A model developed on the basis of these observations suggests that LHCP are transported into the stroma and enter thylakoid membranes in a subsequent step (Reed et al., 1990; Cline et al., 1993). This process is dependent on a developmentally regulated activity (Chitnis et al., 1987) and may describe a pathway operating a t stages of development later than we have studied. ACKNOWLEDGMENT
We wish to thank Dr. Bruce Witmershaus for help with 77 K fluorescence spectroscopy. Received March 21, 1994; accepted June 10, 1994. Copyright Clearance Center: 0032-0889/94/106/0583/08. LITERATURE CITED
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