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Mechanical regulation of photosynthesis in cyanobacteria

Abstract

Photosynthetic organisms regulate their responses to many diverse stimuli in an effort to balance light harvesting with utilizable light energy for carbon fixation and growth (source–sink regulation). This balance is critical to prevent the formation of reactive oxygen species that can lead to cell death. However, investigating the molecular mechanisms that underlie the regulation of photosynthesis in cyanobacteria using ensemble-based measurements remains a challenge due to population heterogeneity. Here, to address this problem, we used long-term quantitative time-lapse fluorescence microscopy, transmission electron microscopy, mathematical modelling and genetic manipulation to visualize and analyse the growth and subcellular dynamics of individual wild-type and mutant cyanobacterial cells over multiple generations. We reveal that mechanical confinement of actively growing Synechococcus sp. PCC 7002 cells leads to the physical disassociation of phycobilisomes and energetic decoupling from the photosynthetic reaction centres. We suggest that the mechanical regulation of photosynthesis is a critical failsafe that prevents cell expansion when light and nutrients are plentiful, but when space is limiting. These results imply that cyanobacteria must convert a fraction of the available light energy into mechanical energy to overcome frictional forces in the environment, providing insight into the regulation of photosynthesis and how microorganisms navigate their physical environment.

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Fig. 1: Quantitative analysis of microcolony formation in Synechococcus sp. PCC 7002.
Fig. 2: Mechanically induced fluorescence emission during microcolony formation.
Fig. 3: The role of phycobilisomes in mechanically induced endogenous fluorescence emission.
Fig. 4: The role of known non-photochemical quenching and mechanical sensing and signalling mechanisms in microcolony formation and endogenous fluorescence emission.
Fig. 5: Modelling the effect of external forces on cell growth.
Fig. 6: Macroscale growth is affected by external forces and photosynthetic source–sink ratio.

Data availability

Data and materials supporting the conclusions of the manuscript are available from the corresponding author on reasonable request. Source data for Figs. 13 and 5 are included with the paper.

Code availability

The code developed to quantify images as well as produce the biomechanical mathematical model was custom generated for this work and is available on GitHub (https://biof-git.colorado.edu/cameron-lab-public/mechanical-regulation-cyanobacteria).

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Acknowledgements

We thank A. Cahill for technical support during setup of the spectral imaging modality, J. Kralj for providing the GCaMP6f plasmid, N. Hill for providing the ∆ccm strain and members of the B. Pfleger laboratory for providing PCC 7002 cells; R. Clark for helpful discussions that improved the manuscript. This research was supported by a RASEI SEED grant (to D.M.B. and J.C.C.). This material is based on work supported by the NSF Postdoctoral Research Fellowship in Biology (grant no. 1711932, to K.A.M.) and in part by an appointment (to S.A.) with the NSF Mathematical Sciences Graduate Internship (MSGI) program sponsored by the National Science Foundation, Division of Mathematical Sciences (DMS). This program is administered by the Oak Ridge Institute for Science and Education (ORISE) through an interagency agreement between the US Department of Energy (DOE) and the NSF. ORISE is managed by ORAU under DOE contract number DE-SC0014664. All opinions expressed in this paper are the authors and do not necessarily reflect the policies and views of NSF, ORAU/ORISE or DOE.

Author information

Authors and Affiliations

Authors

Contributions

J.C.C. conceived the project. J.C.C. and K.A.M. designed the study and analysed the data. K.A.M. generated mutant lines and performed fluorescence microscopy experiments. E.B.J. and J.C.C. performed spot-plating experiments. S.A. and D.M.B. developed the mathematical model. J.C.C. and K.A.M. provided input on model development. J.B.M. performed the electron microscopy. J.W.T. developed the single-cell segmentation and tracking algorithms. J.C.C. and K.A.M. wrote the paper with input from all of the other authors.

Corresponding author

Correspondence to Jeffrey C. Cameron.

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Competing interests

K.A.M., S.A., J.W.T., J.B.M., E.B.J., D.M.B. and J.C.C. are inventors on a patent application filed by the University of Colorado on the basis of this work (patent application no. 16/036,645; filed on 16 July 2018). The authors declare they have no other competing interests.

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Extended data

Extended Data Fig. 1 Representative cell number and endogenous fluorescence traces.

Each panel represents number of cells (black) and mean endogenous fluorescence intensity (emission Cy5 705/72 nm following 640 nm excitation) of all cells (blue) during microcolony formation. Data is from four individual microcolonies, which started with either one or two cells. The red arrow indicates the beginning of fluorescence rise at the 4-cell colony stage.

Extended Data Fig. 2 Transmission electron micrographs of plunge-frozen cyanobacterial microcolonies.

Panels represent en face cross-sections through a Synechococcus sp. PCC 7002 microcolony grown on dialysis tubing on top of 1% agarose A + pads. a, Close-up of cell-cell interfaces in microcolony interior shown in Fig. 2f. b, Close up of microcolony edge. Mechanical deformation of cells is more apparent in the colony interior. Asterisk denotes a cell-lysis event of a highly deformed cell. c, and d, Larger field views of representative microcolonies with distinctive difference between interior and exterior cells. Similar results were observed in two independent experiments.

Extended Data Fig. 3 Analysis of ∆cpc mutant.

a, Absorbance spectra of WT and ∆cpc strains. Consistent results were observed in three independent experiments b, Schematics of WT and ∆cpc genomic regions and location of primers to confirm segregation (not to scale). c, PCR products confirming presence of ∆cpc insert and absence of WT product. Lanes: 1, 1 kb + MW ladder (Bands representing 0.5, 0.1, 1, and 3 kilobases are marked); 2, WT (primers: cpc upstream + GmR); 3, ∆cpc (primers: cpc upstream + GmR); 4, WT (primers: cpc upstream + cpc internal); 5, ∆cpc (primers: cpc upstream + cpc internal). Consistent results were observed in two independent experiments.

Extended Data Fig. 4 Ratio of 651 nm / 684 nm fluorescence emission from single cells during growth on 2% agarose.

Individual cell traces of 20 individual cells that exhibit increased fluorescence while growing on 2% agarose. Y-axis all represent values shown for top left graph. X-axis represent time elapsed from the initial frame of time-lapse imaging.

Extended Data Fig. 5 Ratio of 651 nm / 684 nm emission from single cells during growth on 1% Agarose.

Individual cell traces of 20 individual cells from the 4-cell colony stage. Y-axis all represent values shown for top left graph. X-axis represent time elapsed from the initial frame of time-lapse imaging.

Extended Data Fig. 6 Expanded image of chambers in microfluidic device.

Cells were grown for 15 hr within the microfluidic device. The white line represents the chamber boundary with cells above the line within the 1.1 µm chamber and cells below the line in the 1.3 µm chamber. Emission (Cy5; 705/72 nm) after 640 nm excitation shown. Scale bar = 5 µm. Calibration bar represents intensities between 50-3500 AU (purple to white, respectively) Similar results have been observed in two independent experiments.

Extended Data Fig. 7 Mean endogenous fluorescence emission intensity after nutrient deprivation.

a, Representative images of endogenous fluorescence (Cy5; 705/72 nm) after 640 nm excitation of WT cells grown for 16 hr in shaking flasks in (L to R) full A+ media, A+ media lacking phosphate, and A+ media with 5 % of the standard nitrate concentration. All images shown with the same look up table displaying intensities between 100-4800 AU (purple to white, respectively). Scale bar = 1 µm. Similar results were observed in two independent experiments. b, Box plot of endogenous fluorescence intensities for the population of cells in (a). n = 1961 cells – full media; n = 1180 cells - 0% P; n = 675 cells – 5% N. Boxplots represent same statistics described in Fig. 1e. The mean fluorescence intensity was greater in full media compared to 0% P (corrected P-value = 1.96 × 10-99) and 5% N (corrected P-value = 2.73 × 10-196). The populations are significantly different from each other based on a two sample Kolmogorov-Smirnov test with Bonferroni corrected p-values of < 0.0001 indicated ***.

Supplementary information

Supplementary Information

Supplementary Information, Tables 1 and 2, and references.

Reporting Summary

Suppmentary Video 1

Time-lapse imaging of cells expressing YFP. Left, YFP fluorescence. Middle, endogenous fluorescence with 640 nm excitation (Cy5 emission; 705/72 nm). Right, channel merge. Scale bar, 1 µm. Imaging frequency for Supplementary Videos 1–9 is 10 min. The calibration bar for Supplementary Videos 1 and 4–8 represents intensities of 75–3,500 AU (purple to white, respectively). Similar results were observed in two independent experiments.

Suppmentary Video 2

Time-lapse imaging of cell growth on 0.5% and 2.5% agarose. Left, cells grown on 0.5% agarose. Right, cells grown on 2.5% agarose. Endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) was pseudocoloured to compare differences in autofluorescence between conditions. The calibration bar represents intensities of 60–6,000 AU (purple to white, respectively). Scale bar for Supplementary Videos 2 and 4–9, 5 µm. Similar results were observed in two independent experiments.

Suppmentary Video 3

Time-lapse imaging of ∆cpc cells and control cells expressing YFP. Left, YFP and endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) merge. Right, endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) was pseudocoloured to compare differences in fluorescence between strains. Similar results were observed in two independent experiments.

Suppmentary Video 4

Time-lapse ratiometric imaging of WT cells grown on 2% agarose. WT cells grown on 2% agarose pads. Images were taken using spectral emission collection of 650–720 nm (2.5 nm bandwidth) with 561 nm excitation. Pseudocolouring represents a ratio of 651 nm:685 nm emission from the indicated cell. The calibration bar represents ratios from 0 (blue) to 4 (yellow). Similar results were observed in 20 independent microcolony lineages.

Suppmentary Video 5

Time-lapse imaging of ∆ocp and control cells expressing YFP. Left, YFP and endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) merge. Right, endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) was pseudocoloured to compare differences in fluorescence between strains. Similar results were observed in two independent experiments.

Suppmentary Video 6

Time-lapse imaging of frp CRISPRi and control cells expressing YFP. Left, YFP and endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) merge. Right, endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) was pseudocoloured to compare differences in fluorescence between strains. Similar results were observed in two independent experiments.

Suppmentary Video 7

Time-lapse imaging of the ∆mscL strain. Endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) was pseudocoloured to better visualize changes in fluorescence over time. Similar results were observed in two independent experiments.

Suppmentary Video 8

Time-lapse imaging of the ∆mscS strain. Endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) was pseudocoloured to compare differences in fluorescence between strains. Similar results were observed in two independent experiments.

Suppmentary Video 9

Time-lapse imaging of WT cells (left) and cpcB CRISPRi cells (right) expressing GCaMP, a Ca2+ signalling sensor. Ca2+ signalling was detected by increases in GCaMP6f intensity after excitation at 470 nm (pseudocoloured). The calibration bar represents intensities of 85–850 AU (purple to white, respectively). Similar results were observed in two independent experiments.

Suppmentary Video 10

Mathematical model of microcolony formation. The interior cell colours denote the predicted forces experienced by individual cells: black < 2, 2 < purple < 5, 5 < red < 7, 7 < orange < 10, yellow > 10. The scale bar (right) corresponds to the cell perimeter and denotes relative forces experienced through individual cell–cell interactions. Black = 0 (no force exerted); white = 1 (maximum force).

Suppmentary Video 11

Time-lapse imaging of ∆cpc cells grown on agarose of different percentages. Representative time-lapse-imaging series of endogenous fluorescence from 640 nm excitation (Cy5 emission; 705/72 nm) of ∆cpc cells grown on 0.5% agarose (left) and 1.0% agarose (right). The frame rate and length of time-lapse imaging are identical for both image sets. Similar results were observed in two independent experiments.

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Source Data Fig. 1

Source data for the plots in Figs. 1d,e.

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Source data for the plot in Fig. 2d.

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Source data for the plots in Fig. 3a (inset) and 3c.

Source Data Fig. 5

Source data for the plot in Fig. 5c.

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Moore, K.A., Altus, S., Tay, J.W. et al. Mechanical regulation of photosynthesis in cyanobacteria. Nat Microbiol 5, 757–767 (2020). https://doi.org/10.1038/s41564-020-0684-2

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