Solubilization Method for Isolation of Photosynthetic Mega- and Super-complexes from Conifer Thylakoids

Materials and Reagents

1. Consumables

   1. Conical bottom microcentrifuge tubes, 1.5 ml

   2. Conical bottom centrifuge tubes, 50 ml and 15 ml

   3. Pipet tips (P1000, P200, and P10)




2. Plant material

   P. abies and P. sylvestris L. plant materials collected from naturally grown trees in the Umeå University campus (63.8202° N, 20.3054° E)




3. Reagents

   1. Invitrogen NativePAGE Bis-Tris (3-12% w/v gradient and 1.0 mm thickness, catalog number: BN1001BOX)

   2. Precision Plus Protein Dual Color Standards (SDS-PAGE molecular marker; Bio-Rad, catalog number: 1610374)

   3. SYPRO-ruby staining (ThermoFisher, catalog number: S12001)

   4. APS (Ammonium persulfate, Sigma, catalog number: A3678)

   5. Trizma (Sigma-Aldrich, catalog number: 93362)

   6. Tricine (Sigma-Aldrich, catalog number: T0377)

   7. Sodium Dodecyl Sulphate (SDS) (Sigma-Aldrich, catalog number: 436143)

   8. Serva Coomassie Blue G (SERVA Electrophoresis, catalog number: 17524.02)

   9. Bis-Tris (Sigma-Aldrich, catalog number: 14879)

   10. 6-aminocaproic acid (Sigma-Aldrich, catalog number: A7824)

   11. Glycerol (Sigma-Aldrich, catalog number: G5516)

   12. miracloth (Merk Millipore)

   13. Grinding buffer (B1) (see Recipes)

   14. Shock buffer (B2) (see Recipes)

   15. Storage buffer (B3) (see Recipes)

   16. Aqueous acetone solution (80%) (see Recipes)

   17. Washing buffer 1 (WB1) (see Recipes)

   18. Washing buffer 2 (WB2) (see Recipes)

   19. Washing buffer 3 (WB3) (see Recipes)

   20. Solubilizing BTH buffer (SB) (see Recipes)

   21. n-Dodecyl-α-D-Maltopyranoside (α-DDM) (see Recipes)

   22. Digitonin (see Recipes)

   23. 100 mM α-DDM working solution (see Recipes)

   24. 100 mM digitonin working solution (see Recipes)

   25. Loading buffer (LB) (see Recipes)

   26. Running buffer (RB) (see Recipes)

   27. Soft-staining solution (see Recipes)

   28. Acrylamide solution A (ThermoFischer) (see Recipes)

   29. Acrylamide solution B (ThermoFischer) (see Recipes)

   30. Cathode SDS buffer (CB) (see Recipes)

   31. Anode SDS buffer (AB) (see Recipes)

Equipment

1. Blender (Multiblender Coline, Clas Ohlson)

2. Automatic pipettes (P1000, P200, and P10)

3. Heating block (95°C)

4. Cooled microcentrifuge (Eppendorf, model: 4530)

5. Microcentrifuge tube shaker (Orbit^TM Digital Microtube and Microplate Shaker)

6. Hoefer electrophoretic chamber (model: SE260 Mighty Small II)

7. Bio-Rad electrophoretic chamber (mini)

8. Bio-Rad mini-PAGE casting apparatus

9. Power Supply (Bio-Rad Power pack ultra)

10. pH meter (Bio-Rad)

11. UV-VIS spectrophotometer (Lumina, scanning range 150-800 nm)

Software

1. Chemidoc imaging system (Bio-Rad Image Lab version 6)

Procedure

1. Sample harvesting

   1. Mature needles were cut from branches with a pair of scissors, collected into a glass beaker, and kept on ice (T ≤ 0°C) in the dark.

   2. Needles were harvested on sunny days during June and July from south-facing branches of five different P. sylvestris and P. abies trees growing in the Umeå University campus (63.8202° N, 20.3054° E).

   3. Upon cutting, the samples were collected into a glass beaker (a separate icebox for each biological replicate was used) and placed in an icebox (35-40 cm deep; ¾ filled with ice and ¼ headspace).

   4. After collection of approximately 3-4 small branches (10-15 g of needles), the whole icebox was covered with a double layer of thick-black cloth to ensure darkness in the sunny summer days during the 2-5 min transportation time to the laboratory.

   5. In the laboratory, samples were immediately subjected to isolation in the cold room (T ≤ 4°C) under dim green light. At no point from collecting branches until completion of isolation were samples moved from either icebox/cold room or darkness (see Figure S1 for sampling collection process and working setup).




2. Thylakoid isolation

   Note: This step must be performed in a cold room.

   (Buffers required - B1, B2, and B3; water and buffers were kept in the cold room overnight)

   1. Samples were prepared according to Bag et al. (2020) with slight modifications to maximize protein complex isolation. Briefly, approximately 10-15 g (fresh weight) of needles were washed in ice-cold distilled water; then, 100-150 ml of ice-cold Grinding buffer (B1) was added (1:10 ratio of needles to buffer), and the mixture blended with a high-speed blender and pre-cooled at 4°C for 15 s in five repetitions, with a 30-s gap between each blending step [a high number of blending steps (5-6) at low blending time (15-20 s) and time lapse between each blending steps (25-40 s) are important for isolating intact higher-order complexes].

   2. The blended needles were filtered through two layers of miracloth (Merk Millipore) and centrifuged (4,500 × g for 8 min, at 4°C). The obtained pellet was resuspended in 30-35 ml Shock buffer (B2) and centrifuged (4,500 × g for 8 min, at 4°C).

   3. A subsequent resuspension, this time on Storage buffer (B3), was followed by a two-step centrifugation, first at low speed (200 × g for 2 min, at 4°C, discard pellet), then at higher speed (4,500 × g for 8 min, at 4°C).

   4. The final obtained pellet was resuspended on 800 µl of B3 buffer, aliquoted in small fractions (50-100 µl), and either used immediately or frozen in liquid nitrogen and stored at -80°C for further use. In all steps, the pellet was resuspended with thin hair paintbrushes to prevent excessive mechanical shearing, which might damage the higher-order protein complexes.




3. Chlorophyll concentration determination

   Considering the stoichiometric relationship between Chlorophyll and Chl-binding proteins, the sample concentration is expressed as milligrams of Chls in a milliliter of sample.

   1. Chlorophyll concentration was calculated from 10 µl of samples extracted in 0.99 ml of 80% ice-cold acetone (1 ml final volume) by rigorous vortexing until no clumps could be seen (approximately 2 min; more time may be needed if the sample is very thick).

   2. After vortexing, tubes were centrifuged at 8,000 × g for 5 min and measured according to Porra et al. (1989).




4. Sample preparation, electrophoresis, and isolation of protein complexes in native conditions

   Note: This step must be performed in a cold room.

   Major optimization methods for conifer samples start from this part. From here onwards, steps are denoted chronologically using the same numbers as they appear on the graphical illustration (Figure 1). The volumes and concentrations of the samples, buffers, and detergents only focus on the optimal detergent concentrations that allow maximum separation. Samples, buffers, and detergents required for different tested detergent concentrations are provided in Tables 1 and 2.

   
   

   
   Figure 1. Graphical illustration of the solubilization, gel running, and band identification

   
   

   Table 1. Solubilization with increasing concentrations of α-DDM. The table shows the volumes of solubilizing buffer (SB) and volumes/concentrations of α-DDM used for obtaining the increasing detergent to chlorophyll ratios presented in Figure 2.
   

   *The volumes are referred to the 100 mM α-DDM working solution (see Recipes).

   
   

   Table 2. Solubilization with increasing concentrations of digitonin. The table shows the volumes of SB and volumes/concentrations of digitonin used for obtaining the increasing detergent to chlorophyll ratios presented in Figure 3. Here, a two-steps solubilization was applied: first, 5 mM α-DDM, denoted as a constant +1 µl in the detergent volume column, were added 5 min prior to digitonin, which is added according to the desired concentration (changing volumes in the detergent volume column).

   
   *The volumes are referred to the 100 mM digitonin working stocking solution (see Recipes).

   
   

   1. Washing thylakoid samples

      (Buffers required - WB1, WB2, and WB3)

      Considering the high tannin and polyphenolic contents in conifer needles, separation of native complexes needs several washing steps before solubilization; otherwise, the detergent activity becomes limited. Here, we used three different washing buffers before solubilization. First, 250 µl of WB1 was aliquoted in Eppendorf tubes and a certain volume of thylakoids corresponding to 10 µg of chlorophylls was suspended by mixing by swirling the pipette in the tube (vortexing or pipetting up and down is strictly prohibited since this can destroy protein complexes). After that, the sample was centrifuged (3,000 × g for 2 min, at 4°C) and the supernatant discarded. Then, the same process was repeated twice, once with WB2 and once with WB3.

   2. Thylakoid solubilization

      (Buffers required - SB)

      For thylakoid solubilization, two different approaches are followed based on the detergent used.

      1. First, for α-DDM solubilization, we solubilized the washed thylakoid pellet in 8-9 µl of SB (depending on pellet size, the bigger the pellet, the smaller the volume of SB that should be added) to obtain 10 µl final volume of the resuspended washed thylakoids. In this way, the chlorophyll concentration of the sample becomes 1 mg/ml. After solubilization in SB, 100 mM of n-dodecyl-α-D- maltoside (α-DDM) is added at equal volume (obtaining a final chlorophyll concentration equal to 0.5 mg/ml and a detergent to chlorophyll ratio of 50:1) and solubilized on ice for 30 min at 4°C.

      2. The fractional solubilization process was applied as digitonin solubilization was reported to be unsuccessful in Arabidopsis protocols (Järvi et al., 2011; Rantala et al., 2017). First, thylakoids were solubilized in SB as before to a chlorophyll concentration of 1 mg/ml, then 1 µl of 100 mM n-dodecyl-α-D- maltoside (α-DDM) was added and the sample kept on ice for 5 min. Then, 6 µl of 100 mM digitonin was added and the final volume filled to 20 µl with SB (the final chlorophyll concentration was 0.5 mg/ml and the detergent to chlorophyll ratio 30:1). Gentle shaking was applied to tubes (max 200 rpm in Orbit^TM Digital Microtube and Microplate Shake) and solubilization performed for 25 min at 4°C.

      3. Finally, both solubilized samples were centrifuged (14,000 × g for 25 min, at 4°C) and the obtained supernatant was used as for all analyses presented in this work.

   3. BN-PAGE process

      1. Blue-Native Polyacrylamide Gel Electrophoresis

         (Buffer required - LB and RB)

         BN-PAGE was carried out using 3-12% w/v gradient and 1.0 mm thickness pre-casted 10 well gels. Before loading, samples were prepared at a ratio of 1:10 in loading buffer. A sample volume of 16 µl was loaded, which corresponds to a final concentration of 8 µg/ml Chls in each electrophoretic lane. Before loading the samples on the gel, the precast gels need to be installed on the electrophoretic chamber and the tank needs to be filled with running buffer and cooled down to 4°C. For our samples, we used a sub-zero water bath to keep the temperature of the electrophoretic system continuously below 2°C throughout the running period.

      2. Native electrophoretic conditions

         The run was performed on a Hoefer electrophoretic chamber at 65 V for 14 h (or overnight) at 4°C in the dark. The running buffer had the same composition for both anode and cathode, but before use, the cathode buffer was supplemented with loading buffer at a ratio of 1:1,000.

   4. Post BN-PAGE processes

      1. Soft staining and gel image acquisition

         After the run, the gels were unmounted from the electrophoretic chambers and casting plates and washed in distilled water, fixed, and stained. BN-PAGEs were stained with the soft-staining solution (Figure 2) according to the procedure described by Farci et al. (2017). Gels were imaged on a white-light LED trans-illuminator using a digital camera (Canon EOS RP digital).

         
         

         
         Figure 2. Solubilization of pine (A, B) and spruce (C, D) thylakoids with different concentrations of α-DDM. A and C are RGB images of soft-stained gels taken with a digital camera. B and D are Coomassie-stained images of the same gels.

         
         

      2. 2D Denaturing Polyacrylamide Gel Electrophoresis

         For 2D separation, denaturing Sodium-Dodecyl-Sulfate polyacrylamide gel electrophoresis (SDS-PAGE) was carried out according to Haniewicz et al. (2013). Briefly, the strips from the BN-PAGE were excised and denatured with Rotiload (Roth) at room temperature for 20 min. After denaturation, the strips were placed on top of a 10% denaturing SDS-PAGE (1.5 mm thickness) and sealed with 0.5% (w/v) agarose in cathode buffer. The molecular weight of the complexes isolated by 2D SDS-PAGE was estimated by plotting the retardation factor values (Rf, length of the band migration/length of the dye front) versus the log of the molecular weight of the molecular marker using a polynomial curve fit (second-order polynomial best-fit), according to the manufacturer's instructions. The obtained curve was used for calculating the apparent masses at the equivalent Rf of the 2D SDS-PAGE spots (data not shown). The masses were combined with western-blotting data for the identification of PSI and PSII subunits (see Step 5b).

      3. Denaturing electrophoretic conditions

         The run was performed on a Bio-Rad electrophoretic chamber and was carried out at 75 V for 3 h, at 4°C, in the dark. Freshly made cathode and anode running buffers were used.

   5. Post 2D SDS-PAGE processes

      1. Staining and gel image acquisition

         After the run, the gels were unmounted from the electrophoretic chambers and casting plates and washed in distilled water, fixed, and stained. 2D SDS-PAGEs were stained with SYPRO-ruby staining (Figures 4 and 5) according to the manufacturer’s protocol. Gels were imaged on a white-light LED trans-illuminator using a digital camera (Chemidoc imaging system).

      2. Western blotting analysis

         After 2D SDS-PAGE, proteins were transferred on PVDF membranes and further used for western blotting analysis and identification of PSI and PSII antibodies, as described in Bag et al. (2020).

Data analysis

1. Optimizing concentrations of detergents

   Isolated native thylakoid membranes were solubilized as per the methods in two different detergents n-dodecyl-alpha-D-maltoside (α-DDM) and digitonin. The separation of thylakoid membrane complexes greatly depends on the optimal solubilization of samples (Haniewicz et al., 2015), and the detergent to chlorophyll ratio may vary from species to species (Kouřil et al., 2016 and 2020). For both detergents, we used a series of concentrations, as shown in Figure 2. For α-DDM, we started with the lowest concentration of detergent (0.5% w/vol =10 mM = 10:1 D:Chls) known to release complexes (to our knowledge) in other plant species (Shao et al., 2011), and concentrations were increased up to 70 mM in 10 mM increments (Figure 2). With increasing detergent concentrations, the release of complexes increases and reaches the maximum at 50 mM. It is worth noting that previously it was reported that the use of 40 mM β-DDM releases C₂S₂M₂ as the largest complex, but here, 50 mM α-DDM released even larger complexes from the membrane (Figures 2 and 3), much larger than C₂S₂M₂ complexes found in Arabidopsis. Most likely, these bands could be similar to higher-order super-complexes previously identified in conifers (Kouřil et al., 2020).

     For digitonin solubilization, similar increasing detergent concentrations from 4 mM (0.4% w/vol = 4:1 D:Chls) have been used, but protocols like those used for other species (Järvi et al., 2011) failed to release any complexes at all (Figure S2). We then supplemented digitonin with a very low concentration of α-DDM (5 mM) as this solubilization method is known to result in a better release of higher-order complexes (Wientjes et al., 2013). Moreover, we obtained clearer results with a two-step solubilization (Figure 3), as explained in the methods and previously reported for other species where it was named “differential solubilization” (Haniewicz et al., 2015). From different detergent concentrations, we chose 32 mM of digitonin to achieve maximum release of complexes when used in combination with 0.25 mM α-DDM in a two-steps solubilization process. By this, a total of six higher-order complexes (higher molecular weights than PSII dimer and PSI monomer) could be separated. The bands were named from 1-6 starting from the higher molecular weight (Figure 3E and 3F).

   
   

   
   Figure 3. Solubilization of pine (A, B) and spruce (C, D) thylakoids with different concentrations of digitonin supplemented with 5 mM α-DDM. A and C are RGB images of soft-stained gels taken with a digital camera. B and D are Coomassie-stained images of the same gels. The red boxes marked in B and D are enlarged and depicted at high contrast in the insets E and F, respectively. These show six higher-order complexes (higher molecular weights than PSII dimer and PSI monomer) here named from B1 to B6 starting from the higher molecular weight.




2. Identification of complexes from 2D SDS-PAGE of 1D BN-PAGE strips

   Identification of thylakoid membrane protein complexes separated on native gels have been done in Arabidopsis and Tobacco (Järvi et al., 2011; Haniewicz et al., 2015; Farci et al., 2017). Recently, Grebe et al. (2019) identified proteins from native P. abies complexes released by β-DDM solubilization. We used the information from these maps to identify the protein composition of the complexes from pine (Figure 4C-ii, D-ii) and spruce (Figure 5C-ii, D-ii) and compared these with known Arabidopsis maps by running Arabidopsis thylakoids in parallel (Figure 4 and Figure 5C-i, D-i).

   
   

   
   Figure 4. Identification of possible presence of thylakoid proteins by 2D SDS-PAGE of 1D BN-PAGE strips. Comparison of α-DDM (A) and digitonin solubilization (B) of pine (Psl) and Arabidopsis thaliana (Ath) thylakoid samples. 2D SDS-PAGE of 1D strips (C) cut from α-DDM solubilized Arabidopsis (C-i) and pine (C-ii); 2D SDS-PAGE of 1D strips (D) cut from digitonin solubilized Arabidopsis (D-i) and pine (D-ii).

   
   

   
   Figure 5. Identification of possible presence of thylakoid proteins by 2D SDS-PAGE of 1D BN-PAGE strips. Comparison of α-DDM (A) and digitonin solubilization (B) of spruce (Pab) and Arabidopsis thaliana (Ath) thylakoid samples. 2D SDS-PAGE of 1D strips (C) cut from α-DDM solubilized Arabidopsis (C-i) and spruce (C-ii); 2D SDS-PAGE of 1D strips (D) cut from digitonin solubilized Arabidopsis (D-i) and spruce (D-ii).

   
   

     The 2D SDS-PAGE of α-DDM solubilized BN strips (Figure 4C-ii and Figure 5C-ii) shows that both first and second bands contain mostly PSII proteins, i.e., CP47, CP43, D1, D2, and LHCII (in comparison with band patterns from Grebe et al., 2020). No spots or bands could be assigned to PSI proteins; thus, we conclude that these bands most likely comprised higher-order PSII complexes (either super- or mega- or oligomers of PSII).

     Conversely, in 2D SDS-PAGE of digitonin solubilized BN strips, the first band (Figure 4D-ii and Figure 5D-ii) contains both PSI and PSII proteins, suggesting the presence of any of the three complexes, PSI-LHCI-PSII-LHCII (Rantala et al., 2017), PSII-LHCII, and PSI-LHCI oligomer (Kouřil et al., 2020). The second and third bands were mostly enriched in PSII subunits, while the fourth and fifth bands contain both PSI and PSII subunits. Finally, the last band was mainly represented by PSI core subunits, LHCI, and LHCII, but contained no PSII core subunits, suggesting this band to be either PSI-LHCI or PSI-LHCI-LHCII. This possibility of multiple complexes arises from the fact that, in conifers, there are unique higher-order complexes that are not present in so-called model plants such as Arabidopsis.




3. Future perspectives

   The establishment of this protocol for thylakoid solubilization with non-ionic detergents facilitates further studies of higher-order complexes from conifers, for example, by mass spectroscopy or cryo-EM. This will enable a better understanding of the basis of photosynthetic adaptations in conifers, for example, during the boreal winter.

Recipes

Note: Stock/working solutions and buffers are prepared in Milli-Q water.

1. Grinding buffer (B1)

   50 mM HEPES-KOH pH 7.5

   330 mM sorbitol

   5 mM MgCl₂

   10% (w/v) PEG 6000

   10 mM NaF

   1 mM BSA

   0.65 mM Na-ascorbate

2. Shock buffer (B2)

   50 mM HEPES-KOH pH 7.5

   5 mM MgCl₂

   10 mM NaF

3. Storage buffer (B3)

   50 mM HEPES-KOH pH 7.5

   5 mM MgCl₂

   100 mM sorbitol

   10 mM NaF

4. Aqueous acetone solution

   80% acetone

   20% water

5. Washing buffer 1 (WB1)

   50 mM Bis-Tris HCl (pH 7.0)

   4% sorbitol

   10 mM NaF

   2.5% Pefabloc

6. Washing buffer 2 (WB2)

   40 mM Bis-Tris HCl (pH 7.0)

   5% sorbitol

   10 mM NaF

   2.5% Pefabloc

7. Washing buffer 3 (WB3)

   35 mM Bis-Tris HCl (pH 7.0)

   6% sorbitol

   10 mM NaF

   2.5% Pefabloc

8. Solubilizing BTH buffer (SB)

   25 mM Bis-Tris HCl (pH 7.0)

   20% glycerol

   10 mM NaF

   2.5% Pefabloc

9. α-DDM

   200 mM stock solution in distilled water

10. Digitonin (200 mM stock solution in distilled water)

    Dissolving digitonin requires heating at 92-95°C for 10-12 min.

    If stock solution is frozen for further use, it needs similar heat treatment before it can be used again.

11. 100 mM α-DDM working solution

    25 mM Bis-Tris HCl (pH 7.0)

    20% glycerol

    10 mM NaF

    2.5% Pefabloc

    200 mM α-DDM

    This solution needs to be freshly made every time.

12. 100 mM digitonin working solution

    25 mM Bis-Tris HCl (pH 7.0)

    20% glycerol

    10 mM NaF

    2.5% Pefabloc

    200 mM digitonin

    This solution needs to be freshly made every time.

13. Loading buffer (LB)

    5% serva Coomassie Blue G

    750 mM aminocaproic acid

    50 mM Bis-Tris HCl (pH 7.0)

    35% sucrose

14. Running buffer (RB)

    50 mM BisTris, pH 6.8

    50 mM Tricine

15. Soft-staining solution

    7% acetic acid

    10% ethanol

    83% distilled water

16. Acrylamide 40% (w/v) solution A (ready-to-use) (ThermoFischer)

17. Acrylamide 2% (w/v) solution B (ready-to-use) (ThermoFischer)

18. Cathode SDS buffer (CB)

    0.1 M Tris

    0.1 M Tricine

    0.1% (w/v) SDS

19. Anode SDS buffer (AB)

    0.2 M Tris, pH 8.9

Acknowledgments

This project has received funding from the European Union’s Horizon 2020 research and innovation programme under the Marie Skłodowska-Curie grant agreement No. 675006 to P.B. and S.J. (Umeå University), The Swedish Research council (VR) and the Kempe Foundation to S.J. (Umeå University), and The Carl Tryggers Foundation for Scientific Research CTS 19:324) to D.F and W.P.S. (Umeå University). We would also like to thank Prof. Eva-Mari Aro and Prof. Roman Kouřil for helpful discussions.

Competing interests

The authors declare no conflict or competing interests.

References

1. Albanese, P., Manfredi, M., Meneghesso, A., Marengo, E., Saracco, G., Barber, J., Morosinotto, T. and Pagliano, C. (2016). Dynamic reorganization of photosystem II supercomplexes in response to variations in light intensities. Biochim Biophys Acta 1857(10): 1651-1660.
2. Bag, P., Chukhutsina, V., Zhang, Z., Paul, S., Ivanov, A. G., Shutova, T., Croce, R., Holzwarth, A. R. and Jansson, S. (2020). Direct energy transfer from photosystem II to photosystem I confers winter sustainability in Scots Pine. Nat Commun 11(1): 6388.
3. Barera, S., Pagliano, C., Pape, T., Saracco, G. and Barber, J. (2012). Characterization of PSII-LHCII supercomplexes isolated from pea thylakoid membrane by one-step treatment with alpha- and beta-dodecyl-D-maltoside. Philos Trans R Soc Lond B Biol Sci 367(1608): 3389-3399.
4. Bassi, R. and Dainese, P. (1992). A supramolecular light-harvesting complex from chloroplast photosystem-II membranes. Eur J Biochem 204(1): 317-326.
5. Benson, A. A. and Calvin, M. (1950). Carbon dioxide fixation by green plants. Annu Rev Plant Physio 1(1): 25-42.
6. Bassham, J. A., Benson, A. A. and Calvin, M. (1950). The path of carbon in photosynthesis. J Biol Chem 185(2): 781-787.
7. Chang, C. Y., Brautigam, K., Huner, N. P. A. and Ensminger, I. (2021). Champions of winter survival: cold acclimation and molecular regulation of cold hardiness in evergreen conifers. New Phytol 229(2): 675-691.
8. D'Amici, G. M., Timperio, A. M. and Zolla, L. (2008). Coupling of native liquid phase isoelectrofocusing and blue native polyacrylamide gel electrophoresis: a potent tool for native membrane multiprotein complex separation. J Proteome Res 7(3): 1326-1340.
9. Farci, D., Kirkpatrick, J. and Piano, D. (2017). A new procedure for fast soft staining of BN-PAGEs on photosynthetic complexes. Electrophoresis 38(3-4): 441-446.
10. Ferroni, L., Angeleri, M., Pantaleoni, L., Pagliano, C., Longoni, P., Marsano, F., Aro, E.M., Suorsa, M., Baldisserotto, C., Giovanardi, M., Cella, R. and Pancaldi, S. (2014). Light-dependent reversible phosphorylation of the minor photosystem II antenna Lhcb6 (CP24) occurs in lycophytes. Plant J 77(6): 893-905.
11. Grebe, S., Trotta, A., Bajwa, A. A., Suorsa, M., Gollan, P. J., Jansson, S., Tikkanen, M. and Aro, E. M. (2019). The unique photosynthetic apparatus of Pinaceae: analysis of photosynthetic complexes in Picea abies. J Exp Bot 70(12): 3211-3225.
12. Grebe, S., Trotta, A., Bajwa, A. A., Mancini, I., Bag, P., Jansson, S., Tikkanen, M. and Aro, E. M. (2020). Specific thylakoid protein phosphorylations are prerequisites for overwintering of Norway spruce (Picea abies) photosynthesis. Proc Natl Acad Sci U S A 117(30): 17499-17509.
13. Haniewicz, P., De Sanctis, D., Büchel, C., Schröder, W. P., Loi, M. C., Kieselbach, T., Bochtler, M. and Piano, D. (2013). Isolation of monomeric photosystem II that retains the subunit PsbS. Photosynth Res 118(3): 199-207.
14. Haniewicz, P., Floris, D., Farci, D., Kirkpatrick, J., Loi, M. C., Buchel, C., Bochtler, M. and Piano, D. (2015). Isolation of Plant Photosystem II Complexes by Fractional Solubilization. Front Plant Sci 6: 1100.
15. Hill, R. (1937). Oxygen evolved by isolated chloroplasts. Nature 139(3525): 881-882.
16. Hill, R. (1939). Oxygen produced by isolated chloroplasts. Proc R Soc London Ser. B-Biological Sci 127(847): 192-210.
17. Huzisige, H. and Ke, B. (1993). Dynamics of the history of photosynthesis research. Photosynth Res 38(2): 185-209.
18. Iwai, M., Grob, P., Iavarone, A. T., Nogales, E. and Niyogi, K. K. (2018). A unique supramolecular organization of photosystem I in the moss Physcomitrella patens. Nat Plants 4(11): 904-909.
19. Järvi, S., Suorsa, M., Paakkarinen, V. and Aro, E. M. (2011). Optimized native gel systems for separation of thylakoid protein complexes: novel super- and mega-complexes. Biochem J 439(2): 207-214.
20. Kouřil, R., Nosek, L., Bartos, J., Boekema, E. J. and Ilik, P. (2016). Evolutionary loss of light-harvesting proteins Lhcb6 and Lhcb3 in major land plant groups--break-up of current dogma. New Phytol 210(3): 808-814.
21. Kouřil, R., Nosek, L., Opatikova, M., Arshad, R., Semchonok, D. A., Chamrad, I., Lenobel, R., Boekema, E. J. and Ilik, P. (2020). Unique organization of photosystem II supercomplexes and megacomplexes in Norway spruce. Plant J 104(1): 215-225.
22. Kügler, M., Jänsch, L., Kruft, V., Schmitz, U. K. and Braun, H. P. (1997). Analysis of the chloroplast protein complexes by blue-native polyacrylamide gel electrophoresis (BN-PAGE). Photosynth Res 53(1): 35-44.
23. Nystedt, B., Street, N. R., Wetterbom, A., Zuccolo, A., Lin, Y. C., Scofield, D. G., et al. (2013). The Norway spruce genome sequence and conifer genome evolution. Nature 497(7451): 579-584.
24. Porra, R. J., Thompson, W. A. and Kriedemann, P..E. (1989). Determination of accurate extinction coefficients and simultaneous equations for assaying chlorophylls a and b extracted with four different solvents: verification of the concentration of chlorophyll standards by atomic absorption spectroscopy. Biochim Biophys Acta 975 (3): 384-394.
25. Porter, G. N. (1950). Flash photolysis and spectroscopy. A new method for the study of free radical reactions. Proc R Soc London Ser A Math Phys Sci 200(1061): 284-300.
26. Rantala, M., Tikkanen, M. and Aro, E. M. (2017). Proteomic characterization of hierarchical megacomplex formation in Arabidopsis thylakoid membrane. Plant J 92(5): 951-962.
27. Romanowska, E., Kargul, J., Powikrowska, M., Finazzi, G., Nield, J., Drozak, A. and Pokorska, B. (2008). Structural organization of photosynthetic apparatus in agranal chloroplasts of maize. J Biol Chem 283(38):26037-26046.
28. Schagger, H. and von Jagow, G. (1991). Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal Biochem 199(2): 223-231.
29. Shao, J., Zhang, Y., Yu, J., Guo, L. and Ding, Y. (2011). Isolation of thylakoid membrane complexes from rice by a new double-strips BN/SDS-PAGE and bioinformatics prediction of stromal ridge subunits interaction. PLoS One 6(5): e20342.
30. Shen, J., Williams-Carrier, R. and Barkan, A. (2017). PSA3, a protein on the stromal face of the thylakoid membrane, promotes photosystem I accumulation in cooperation with the assembly factor PYG7. Plant Physiol 174(3):1850-1862.
31. Wientjes, E., Drop, B., Kouřil, R., Boekema, E. J. and Croce, R. (2013). During state 1 to state 2 transition in Arabidopsis thaliana, the photosystem II supercomplex gets phosphorylated but does not disassemble. J Biol Chem 288(46): 32821-32826.
32. Wood, W. H. J., MacGregor-Chatwin, C., Barnett, S. F. H., Mayneord, G. E., Huang, X., Hobbs, J. K., Hunter, C. N. and Johnson, M. P. (2018). Dynamic thylakoid stacking regulates the balance between linear and cyclic photosynthetic electron transfer. Nat Plants 4(2): 116-127.