The Utilization of Varying Wavelengths of Light by Cultured and Environmental Cyanobacteria

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The utilization of varying wavelengths of light by cultured and environmental cyanobacteria

James R. Henriksen Microbial Diversity Course 2010, MBL

Abstract

Photoautotrophy is the foundation of all ecosystems. In order to explore the vast diversity of mechanisms used by bacteria to capture light energy, two methods were developed: a high-throughput method for physiological investigations and selective enrichments of speciﬁc organisms based on varying wavelength and intensity of light, and a culture independent, microscopic method based on autoﬂuorescence for probing the light utilization and components of microbial photosystems. These were applied in an investigation of cyanobacterial light harvesting apparatus in cultured isolates and environmental samples that provided insights into cellular variation and life cycle in isolated cyanobacterial cultures and the ecological physiology and selection in a “Green Berry” cyanobacteria-dominated microbial consortium.

Background

Microorganisms in the environment, particularly prokaryotes, are major drivers in all biogeochemical cycles, and are critical for providing ecosystem functions (Madigan 2008). The vast diversity of physiologies and characteristics are the very fabric of life on this planet, and may increase the stability of the biosphere. An understanding of the the vectors in Hutchinsonian niche space that lead to specalisation and evolutionary pressures, and an understanding of the mechanisms of utilising these diverse resources are the major pursuits of environmental microbiology (Holt 2009). One such vector at the base of all foodchains are the wavelength and amount of light present for photosynthesis. Oxagenic photosynthesis drives almost all energy cycling and carbon fixation on the planet. Even the 0.8% of global carbon fixation that is carried out by anoxic autotrophy is dependent on oxagenic photosynthesis, as the oxidant and reductants in all modern food webs, even deep sea hydrothermal vents, originate from oxagenic photosynthesis (Raven 2009). Oxagenic photosynthesis evolved only in the cyanobacterial lineage, probably from a horizontal gene transfer that resulted in the combination of the photosystems from green and purple bacteria (Madigan 2008). Cyanobacteria are responsible for 50% of the global primary productivity and the majority of carbon that is rapidly cycled, with the remainder being provided by the cyanobacterial endosymbionts (chloroplasts) of eukaryotes (Phillips 2009). The major components needed for the growth of oxygenic diazotrophic photoautotrophs are available in abundance. These organisms require only light, water, common gases found in the atmosphere, and trace elements. It is instructive to consider the amounts of these components found in nature. All values for the following are calculated from information in the Bionumbers database (Phillips 2009) and the CRC Handbook of Chemistry and Physics (Haynes 2010). Water is the most common electron donor available that is accessible to life, at concentrations of ∼55 M (the molarity of pure elemental water) over the majority of the plants surface. Dinitrogen gas is the most abundant atmospheric gas, present at 3.5 moles/L air at sea level, but only at ∼45 µM in seawater, due to it’s poor solubility. However, this concentration is enough to meet the nitrogen demand of ∼107 bacterial cells/ml of seawater, even without taking into account a net flux from the atmosphere. The carbon dioxide that is the sole source of carbon for autotrophs is present at 17mmols/L in the atmosphere at sealevel, and at average of ∼2mM in seawater, sufficient for ∼108 bacterial cells/ml of seawater. Trace elements needed by oxigenic diazotrophic photoautotrophs are only needed at low levels, but may be limiting in parts of the ocean where dust inputs are low. 1 Light, the final requirement of oxigenic diazotrophic photoautotrophs, and their sole source of energy, is abundant at the surface of the planet. If an organism has the machinery to access it, light is an excellent source of energy. The energy available in light varies with its wavelength (sometimes termed quality, measured in nm or in inverse frequency in Hz) and the number of photons (intensity or quantity measured in mols of photons, or Einsteins, per second per area). Light with wavelengths of 400 to 1200nm provides -350 to -100 kJ/mol (Overmann 2000), and in direct sunlight photons across the spectrum are provided at 2000 µEinsteins/m2/sec, close to the maximum flux used in photosynthesis (Phillips 2009). For a cell with a photosynthetic cross section of 10µm, this would correspond to 1010 photons per sec. For comparison, E. coli growing aerobically with a doubling time of 20 minutes use glucose at -2870 kJ/mol at a turnover rate of 106 mol/sec (Phillips 2009). Therefore, light provides less energy per electron transferred, but is available at a high rate. The wavelength and intensity of light available varies in the environment (particularly with depth, as shorter wavelengths of light penetrate deeper in bodies of water). The wavelength and intensity of light that can be utilised or tolerated between organisms also varies. The the minimum energy and intensity of light thought necessary to support life are 117 KJ/mol (corresponding to a wavelength of 1020nm, Overmann 2000) and 10-5 µE/m2/s (Beatty 2005) respectively. The maximum energy of light is assumed to be limited by the toxisity of far UV light. The major groups of phototrophs (PSB, PNS, GNS, cyanobacteria, green and red algae, and diatoms) utilize different spectra of light (termed the action spectra) and use different photosynthetic apparatus. This may be based on the molecular characteristics of light absorption water, as well as light characteristics of different bodies of water (Stomp 2007). Cyanobacteria are unique among oxygenic photoautotrophs in their ability to capture light across the visible range, including the green light that is not utilised by eukaryotic phototrophs. Cyanobacteria, like many phototrophic groups, contain organisms that have specific characteristic wavelengths of light that they utilize. However cyanobacteria are unique in that some organisms can change the wavelength of light that can be captured by producing different phycobiliproteins, a process termed complementary chromatic adaptation. Cyanobacteria use a diverse set of phycobiliproteins in various combinations in a phycobilisome antenna complex. All cyanobacteria produce the phycobiliproteins allophycocyanin and phycocyanin, while some also have phycoerythrin or phycoerythrocyanin. These proteins contain chromophore tetrapyrroles known as phycobilins that along with the protein and modifications to the structure determine the wavelength of light captured. Different phycobiliproteins contain different combinations of phycobilins (see Table 1). All cyanobacteria produce phycobilins phycocyanobilin and phycoerythrobilin, and some can produce phycobiliviolin (also known as phycoviolobilin or cryptoviolin) or phycourobilin (see Table 1). Some cyanobacteria constitutively express all phycobiliproteins they can produce, while others can regulate some or all of their phycobiliproteins. A number of factors can change the amounts, ratios and types phycobiliproteins and chlorophyll reaction centers. Of these the best characterised in complementary chromatic adaptation species is the available amount and wavelength of light. Other factors that might influence the light absorbed include the characteristics of the organisms, nitrogen and CO2 availability, and cellular stress. Cyanobacterial photosynthesis is composed of a z scheme where a photon of light excites antenna pigments in the phycobilisomes. If the organisms is producing them, lower wavelength (higher energy) photons are absorbed by other phycobiliproteins and their energy is transferred to phycocyanin and then to allophycocyanin. The energy from the phycobilisome is passed PSII where charge separation traps the energy in electron potential and splits water to O2. This energy is then passed to through a electron transport chain (generating ATP) to PSI, which, when excited by another photon, passes the energy along another electron transport chain to either the chain of PSII in cyclic electron flow or to form NADH in a non-cyclic man- ner. Besides this two-photosystem system, all cyanobacteria can carry out PSI-based sulfate oxidation by degrading PSII and the phycobiliproteins. There are some observations that suggest that phycobiliproteins may in some cases pass energy to PSI, but this is controversial. The excitation peaks of the phycobiliproteins are broad and often show significant tailing and variation between sub-types in different organisms. The characteristic maxima of the spectral curves of the major components of the Cyanobacterial photosynthetic machinery are listed in Table 1. The florescent emission spectra of any compound is characteristic for a molecule. If the absorbed light energy causes fluorescence, the absorbance peak is also the fluorescent excitation maxima. If energy is passed from one component to another (usually from phycobiliproteins to chlorophil), the observed fluorescence emission will be that of 2 Table 1: Absorbance and fluorescence values for cyanobacterial photosystems. Values are from whole cells and peptide-bound billins, not extracts or pure compounds. Note that the exact shape of the absorbance and the specific fluorescence maxima can be determined, and are diagnostic of a compound in a specific intracellular environment. Ranges are compiled from Overman 2010, Fay 1987, Platt 1986 and Glazer 1988.

Florescent Photopigment Components Absorbance (nm) emission (nm) R-phycoerythrin phycoerythrobilin, phycourobilin 455-575 538-581 B-phycoerythrin phycoerythrobilin, phycourobilin 475-575 538-581 C-phycoerythrin phycoerythrobilin 494–575 538-581 phycoerythrocyanin phycocyanobilin, phycobiliviolin 568-590 568-625 R-phycocyanin phycocyanobilin, phycoerythrobilin 533-615 640-650 C-phycocyanin phycocyanobilin 615–630 617-645 allophycocyanin phycocyanobilin 650–655 650-680 chlorophyll a chlorophil a 400, 660-700 680-720

chlorophil, while the excitation maxima will be the absorption maxima of the initial absorbing component. The transfer of energy between components is very efficient, with more than 98% of the energy being captured delivered to chlorophil before it can be re-emited as heat or fluorescence. Energy is usually only lost at the chlorophil molecule, causing fluorescence. The quantum yield of the photosynthetic reaction complex is high, and the autoflourecence seen from these systems are within the range for fluorophores used in fluorescent microscopy. The pattern of excitation and emission is characteristic of the presence of the various parts of the components in the photochain. Cyanobacteria occur in many environments, and are responsible for large amounts of primary productivity in the open ocean. One coastal ecosystem where macroscopic aggregates of cyanobacteria are visually dominant is in pools found in Sippiwisit Marsh that are filled with small (approximately 1-5mm) spherical “green berries”. For previous work by a Microbial Diversity Course student on the macroscale dark green cyanobacterial aggregates, see Gentile 2002.

Materials and Methods Cultures The unicellular marine cyanobacteria Cyanothece ATCC51142 and Synechococcus PCC7335, and the unicellular freshwater, low-light adapted Gloeothece PCC7109 were kindly provided by John Waterbury. Anabena PCC7120 was kindly provided by Bob Haselkorn.

Environmental samples Macroscale dark green cyanobacterial aggregates (“green berries”) were kindly collected by Parris Humphrey, Ulli Jaekel, and Lizzy Wilibanks from pools in the Lesser Sippiwisit Marsh, a salt-water marsh north of Woods Hole, Massachusetts where they co-occur with purple bacterial aggregates. Three berries were homogenised with a microcentrifuge pestle and resuspended in media by vortexing.

Culture conditions Cultures were grown on SN or BN media (Waterbury 2006) for marine and freshwater strains respectively at 30C inoculated from liquid stock cultures. All pre-cultures were grown for one week under broad-spectrum fluorescent lights at 75 µEinsteins/m2/s unless otherwise noted. All strains were checked for nitrogen fixation in media lacking a nitrogen source, and examined microscopically for morphology. 3 Table 2: LED specifications

Luminous Forward Unit Wavelength Flux at Current Viewing Supplier and Color / Name Voltage Price (nm) Current (A) Angle Part # (Vf) (US$) (mlm) All Electron- UV 380 100mA 3.5 ∼45◦ 0.5 ics LED-910 Digi-Key Blue 470 495 20mA 3.4 85◦ 0.52 365-1201-ND Digi-Key Green 525 1110 20mA 3.4 85◦ 0.58 365-1202-ND Digi-Key Amber-Yellow 592 278 20mA 2 30◦ 0.25 160-1503-ND Digi-Key Orange-Yellow 605 278 20mA 2 30◦ 0.29 160-1502-ND Digi-Key Orange-Red 615 278 20mA 2 30◦ 0.29 160-1501-ND Digi-Key Red 623 941 20mA 2.2 85◦ 0.3 365-1203-ND not re- not re- Digi-Key IR 850 1.5 26◦ 0.3 ported ported 475-1460-ND Digi-Key C535A- 900K White broad 3750 20mA 3.2 110◦ WJN- 0.43 CS0V0151- ND

Multi-spectral LED culturing plates electronics A led lightbox was prepared with cheap commercially available 5mm through-hole LEDs arrayed in the wells of a black polystyrene 96 well plate (see Table 2). Each row consisted of one of each LED type wired in series so as to provide a constant voltage drop. These were connected in parallel with a 100W high wattage resister to limit current spikes and driven by a regulated power supply in series with a digital multimeter to measure current. At a current through the LEDs of 150mA (slightly less than the test current for the LEDs of 20mA times 8 parallel LED circuits), the voltage across the LED array was close to the 19.86 predicted by Ohm’s law and the typical forward voltage of the LEDs reported on their specification sheets. The power supply was set so as to be voltage limiting, but any increase in current would cause it to become current limited. These precautions should prevent any problems with burning out of LEDs. The LED wavelengths (from specification sheets and confirmed with a handheld diffraction grating spectrometer) and intensities are shown in Table 3. Typical LEDs have spectral half-max widths of 5-10nm.

Multi-spectral LED culturing plates Stacks of sterile clear-bottomed and clear-lidded, black-walled polystyrene 96 well plates were placed on top of the lightbox, such such that each well received light from one LED. Stack of plates of 200ul of cultures were interspaced with 96 well 50% neutral density plates. The neutral density plates were also clear-bottomed 96 well plates with 200 µl of a dilution of 0.12g activated charcoal in 50% saturated sucrose. Linear and logarithmic dilutions of this suspension were made on a 96 well plate and measured with a spectrophotometer to determine the dilution that archived 50% transmittance.

4 Table 3: LED spectral characteristics

Calculated Quantum Approximate Wavelength photon Pyranometer sensor Color / Name Flux (nm) energy measurements measure- (µE/m2/s) (KJ/E) ments UV 380 288.42 3.9 101.25 Blue 470 233.19 43.35 550 100 Green 525 208.76 19.55 450 150 Amber-Yellow 592 185.14 2.7 200 100 Orange-Yellow 605 181.16 3.3 300 75 Orange-Red 615 178.21 8.1 200 210 Red 623 175.92 15.3 450 180 IR 850 128.94 1500 900K White broad 150 60

96 well light culturing High throughput LED culturing chambers were used to expose multiple cultures (up to 8 per plate) to each of the different wavelengths of light, at different intensities (using the charcoal-glycerol neutral density filters). Wells in three 96 well plates were filled with 100ul of an appropriate medium (SN or BN). All wells in a row were inoculated with one of the pure cultures or consortia, or left uninoculated as a sterility control. Since the different light conditions were arrayed in columns on the plate, each inocula was exposed to one of the 10 different wavelengths, at three intensities (100%, 50%, 25% of values listed in Table 3), as well as multiple dark controls. During use the LED light array was operated with a timer for 16h light / 8h dark and the cultures kept at 30in an incubator Absorbance and fluorescence were measured daily.

Absorbance and fluorescence A Molecular devices M5 plate reading flouromiter was used for all well absorbance (at 600nm and the Chlorophyll absorbance maximum) and fluorescence measurements. For fluorescence, the following excitation wavelengths were used, corresponding to the LED wavelengths: 380, 470, 525, 592, 605, 615, 623, and 850nm. For each, the closest filter set was used, and all emission scans were started at least 50nm from the excitation source to limit ramen scattering and other spurious effects. An automated program was used to collect absorbance and each of the fluorescent emission spectral scans. Please contact the author for copies of the program that automates the multi-spectral scanning of the 96 well plates.

Light intensity Intensities were measured by a Li-Cor photometer (model LI-185-B) with a PAR photosensor (LI200SB pyranometer) or a quantum sensor (LI 190S). The meters were not corrected for spectral sensitivity, but were calibrated at at 150 mA µE/m2/s.

Microscopic slide preparation For microscopic spectral determination, 10ul samples scraped from the bottom of a well were placed on agar coated slides, covered with a coverslip, and sealed with clear ﬁngernail polish. The agar slides were prepared by coating a clean glass slide with hot 1% agar dissolved in water and allowing them to dry for 24h.

Spectral confocal scanning laser microscopy (spectral-CSLM) A Ziess 710 spectral confocal scanning laser inverted microscope was used for all autoﬂourecence spectral microscopy. For each 2nm bandwidth excitation laser (405, 458, 488, 514, 561, 594, and 633nm), the percent 5 Table 4: Table 4: 710 lasers and settings

Laser (nm) Laser power (mW) %T setting 405 30 6.67 458 2.63 76 488 13.16 15.2 514 6.58 30.4 561 15 13.33 594 2 100 633 5 40

transmittance was adjusted to provide a maximum laser output of 2mW (see Table 4). The PMT in the 710 varies linearly from 30% quantum eﬃciency at 400nm, to 10% at 700, and the detectors are tuned to deliver a near-linear response across the full wavelength (Zeiss representative, personal communication). The automated capabilities of Zeiss ZEN software were used to automate the multi-spectral imaging and scans that collected z-stack, tiled image, or timelapse data. Please contact the author for copies of the settings ﬁles, program, and scripts that automate the multi-spectral scanning.

Data analysis and visualisation A script in the R programming language (R Core Development Team 2010, Gentlemen 2008) was used to extract data from comma separated values ﬁles exported from the M9 and Zeiss ZEN software, and generate tiled plots using a grammar of graphics-based package (Wikinson 1999, Wickam 2009). Please contact the author for copies of script.

Results and Discussion

While autoflourecence, or the fluorescence from phototropic microorganisms, is usually viewed as a nuisance by microscopists wishing to observe their own introduced fluorophore markers (such as FISH), there is information available in the excitation and emission spectra of this autoflourecence. While photobleaching might indicate maximum usable fluxes, the most accessible information is the wavelength of light that causes fluorescence, and the wavelength of the emitted fluorescence In particular, if light energy is absorbed by phycobiliproteins and emitted by chlorophyll, the types of phycobiliproteins and their coupling to the photosystem can be deduced. The functioning is much like a tightly coupled, multiple- fluorophore FRET using the coupled antenna and photosystem chain instead of two fluorescent molecules in close proximity. This is the basis of the autoflourecence spectral microscopy technique developed in this work. After developing this technique, two papers with similar methods were found (Polerecky 2009, Roldán2004). To the author’s knowledge this work is the first to utilise this method in cyanobacterial macro-consortia, the first to utilise CSLM and multiple wavelength lasers, and the first to couple it to enrichment based on wavelength of light. Four pure cultures and a consortium (Figure 1) were grown under three intensities and eight monochro- matic LEDs, as well as broad-spectrum white LED and dark controls. Growth densities were followed by absorbance for 7 days (example absorbance data shown in Figure 2). The culture or enrichment bulk fluorescence was measured in each 96 well plate. An example of the fluorescence data is shown in Figure 3. While little or no growth was seen for many of the organisms under many of the light regimes, some culture increased in absorbance under some light regimes. Also note that for cultures with low growth, the bulk fluorescence is not informative. Stock cultures grown for more than one month with diffuse sunlight were sampled and their photosytems examined using spectral-CSLM analysis of their photosystems autoflourecence. The different fluorescence spectra acquired are displayed in Figures 4, 5, 6 and 7 and initial results are discussed below. While initial data was collected for the pure cultures grown under different wavelengths in the multi-spectral LED culturing plates, a lack of several doubling in many of the cultures due to insufficient growth time prevents 6 the interpretation of the autoflourecence state of the cells. Instead, the results interpreted below will focus on the death phase stock cultures and the changes to the “Green Berries” consortia enriched with various wavelengths on multi-spectral LED culturing plates. Cyanothece cells displayed the most restricted excitation spectra (Figure 6), with typical 650nm maximum chlorophil fluorescence observed only with 561, 594, and 633nm excitation. This indicates that under these conditions the predominant phycobiliproteins present are not the blue-shifted phycoerythrins or phycoerythrocyanin, as nothing with their absorbance pattern transferred energy to chlorophyll, nor were their uncoupled fluorescence observed. All Cyanothece cells gave similar autoflourecence emission / excitation spectra. This data corresponds well to the 96 well bulk culture fluorescence. In the Anabena cultures, there was a strong coupling of 380nm excitation to 675nm maximal fluorescence, probably from a the lower absorbance band of chlorophil. There was a lower emission from across all the longer laser wavelengths. Heterocysts (region 23 in Figure 7) gave reduced fluorescence, and in other cultures decoupling of phycobiliproteins was observed during the dismantling of photosystem I during heterocyst development (data not shown). Gloeothece cells displayed two different autoflourecence emission / excitation spectra. Most cells in the culture (Figure 4), like actively growing cultures (data not shown) had a fluorescence maxima at either 650 or 725nm or both, depending on the excitation wavelength. This indicates multiple fluorophores, either coupled to different chlorophylls or longer-wavelength emitting phycobiliprotein that is not coupled to any chlorophil. Occasional cells were observed that had fewer intracellular vacuoles and a lower density in the DIC micrograph which corresponded to a fainter fluorescence with less fluorescence shift (Regions 16 to 20 in Figure 4). This was probably due to the degradation of portions of the photosynthetic apparatus. Likewise, there was a presumed degradation of some Synechococcus cells. While preliminary, this analysis of the spectral-CSLM analysis of their photosystems autoflourecence indicates that differences in single-cell autoflourecence can reveal details of photosystem function. For instance, Gloeothece grown under these conditions would seem to be a photo-generalist, while Cyanothece seems to utilize a more restricted wavelength of photons. In addition, spectral-CSLM analysis of autoflourecence shows a decoupling and degradation of photosystem components during the death phase of cultures. The homogenised “Green Berry” consortia grown in wells on the highest light exposure plate and exposed to 4 different light conditions are displayed in Figures 8, 9, 10, 11, and 12. The spectral patterns and cellular abundances found in the white LED enrichments and initial micrographs are most smiler to the cultures enriched at 470nm LED (Figure 8 and 9, respectively). The dominant Gloeothece-morphology organisms (Gentile 2002) enriched with all light treatments were particularly prevalent at 470nm LED light enrichment (Regions 2 to 4 in Figures 8, 10 11, 12 and 13 as well as Regions 2 to 16 in Figure 9). These Gloeothece- morphology organisms grown at 470nm display a similar pattern to the pure culture Gloeothece grown in sunlight, in that they have a strong emission maximum at 725nm with excitation at 594 and 633nm, and a shorter wavelength emission with broad excitation. However, the exact emission spectra were slightly shifted at the middle frequencies. In all other enrichments, the show a different pattern, fluorescing less below 514nm. Filamentous phototrophs were observed as minor organisms is lower-wavelength enrichments (Regions 8-10 in Figure 8, Regions 11 to 13 in Figure 10 and Regions 8-13 in Figure 11) with two strikingly different emission fluorescence spectra (compare Regions 8 to 10 and 11 to 13 on Figure 11). The Blue-shifted fluorescence seen in many of the cultures is very unusual, and does not correspond to known chlorophyll emission spectra. Diatoms with morphologies similar to stramenopiles and dinophyceae were found in many micrograph of the Green Berries, and all had the expected patterns of fluorescence (see Regions 5 to 7 in Figure 8, Regions 17-25 in Figure 9, regions 5 to 7 in Figure 10, and Regions 5 to 10 in Figure 12). They were particularly abundant in enrichments at 561, 594, and 633 nm (Figure 12 and data not shown). Enrichments based on the wavelength of light provided by multi-spectral LED 96 well culturing plates impacted the morphological makeup of the consortia, with diatoms out-competing the cyanobacteria at higher wavelength. Using the autoflourecence of naturally coupled photosystems with spectral CSLM is a powerful technique to investigate both single-cell heterogeneity, development, and ageing in pure culture, and the photosystems and light utilization of not-yet cultured organisms. Green Berries contain organisms that can utilize diverse wavelengths of light, and the dominant Gloeothece-morphology organisms may photoadapt at 470nm. Interesting diversity, even within one filamentous morphology in the same light enrichment was seen, and potentially novel photosystem were observed using this technique.

7 Future work

High throughput isolation of organisms with varying light conditions is possible with the Multi-spectral LED culturing plates developed for this work. Sequencing of communities exposed to different LED light treatments would allow more in-depth measure of the enrichment effects, and potentially tie organisms to observed spectral patterns. Investigations of complex photosynthetic communities could be pursued with the autoflourecence spectral microscopy technique developed here. Further conformation, as well as extractions of pigments (with both water and acetone) could lead to more precise understanding of uncultivated organisms’ photosystems, particularly photoadaptation with phycobilins and novel chlorophylls. Microrespirometry of wells lit with a single wavelength LED would connect the absorbance measurements to the efficiency of energy capture at various wavelengths. Pigments that are not involved in photosystems, such as that produces by Pseudomonads in biofilms could also be used with the autoflourecence spectral microscopy technique to act as a spectral fingerprint. A full calibration and propagation of spectral sensitivities with this technique could also provide a quantification of light utilization. Extending this technique with time resolved and polarised florescence could provide data on the the dynamics and the bound state of the fluorophores respectively.

Acknowledgements

I would like to acknowledge Steven Zinder, Dan Buckly, Bill Metcalf and all of my classmates and instructors at the 2010 Microbial Diversity Course at MBL for many hours of discussion and friendship. These weeks have been the most intense, interesting, and exhausting of my academic life. Thanks for the great companionship and great times. I wish to thank John Waterbury and Bob Haselcorn, and Team Berry (Parris, Ulli, and Lizzy) for cultures and advice. I would like to thank Rudi Rottenfusser, Chris Reikin and Alex Valm for fruitful discussions about quantitative CSLM. In addition, I wish to thank my mentor, Yosko Fujita, for her generous help and support in attending this course. This work was supported by grants from Idaho National Laboratory Division Initiative and Biological Systems Departmental Grants, The Gordon & Betty Moore Foundation, and the National Aeronautic and Space Administration.

References

Beatty, J.T. et al., 2005. An obligately photosynthetic bacterial anaerobe from a deep-sea hydrothermal vent. Proceedings of the National Academy of Sciences of the United States of America, 102(26), 9306 -9310. Fay & Wyman, 1987. Acclimation to the natural light climate. In The Cyanobacteria. Amsterdam: Elsevier Science Publishers, p. 348. Gentile, M., 2002. The Green Berries. MBL Final Project. Gentleman, R., 2009. R programming for bioinformatics, Chapman & Hall/CRC. Glazer, A.N., 1988. Phycobiliproteins. Methods in enzymology, 167, 291. Glazer, A.N., 1987. Phycobilisomes: Assembly and attachment. In: The Cyanobacteria (P. Fay and C. Van Baalen, eds.), Elsevier, Amsterdam. Haynes, D.R., 2010. CRC handbook of chemistry and physics, CRC press. Holt, R.D., 2009. Colloquium Papers: Bringing the Hutchinsonian niche into the 21st century: Ecological and evolutionary perspectives. Proceedings of the National Academy of Sciences, 106(Supplement 2), 19659-19665. Madigan, M.T. et al., 2008. Brock Biology of Microorganisms 12th ed., Benjamin Cummings. Overmann, J. & Garcia-Pichel, F., 2006. The phototrophic way of life. In The Prokaryotes, 32–85. Dworkin M., editor, New York: Springer. Phillips, R. & Milo, R., 2009. A feeling for the numbers in biology. Proceedings of the National Academy of Sciences, 106(51), 21465 -21471. Platt, T. & Li, W.K.W., Photosynthetic picoplankton. Canadian Bulletin of Fisheries and Aquatic Sciences, 214, 582. 8 Raven, J.A., 2009. Contributions of anoxygenic and oxygenic phototrophy and chemolithotrophy to carbon and oxygen fluxes in aquatic environments. Aquatic Microbial Ecology, 56, 177–192. Stomp, M. et al., 2007. Colorful niches of phototrophic microorganisms shaped by vibrations of the water molecule. ISME J, 1(4), 271-282. Team, R., 2008. R: A language and environment for statistical computing. R Foundation for Statistical Computing Vienna Austria ISBN, 3(10). Waterbury, J.B., 2006. The cyanobacteria: isolation, purification, and identification. The Prokaryotes, 1053–1073. Wickham, H., 2009. ggplot2: elegant graphics for data analysis, Springer-Verlag New York Inc. Wilkinson, L., 2005. The grammar of graphics, Springer Verlag.

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Figure 2: OD at multiple wavelengths (see legend) indicating growth over time (x axis) for a blank and 8 strains (upper grey tabs, JW=original stock cultures >1month old, HL=pre- cultures grown under high intensity florescent lights), grown at three light intensity levels (right grey tabs). Figure 3: Fluorescence of culture wells in 96well plate. Each of the 66 tiles in this array represents measurements on one well. The grey tabs along the top indicate the light exposure (0=dark, 1=white LED, and NA=empty well blank, all others are wavelength in nm), the abbreviations along the grey boxes at the right indicate the organism (blank = uninoculated, Ana=Anabaena, Cya=Cyanothece, GB=green berries, Glo=Gloeothece, Syn=Synechococcus). Each tile displays the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). The intensity value of red color indicates the relative florescent units. Figure 4: Gloeothece cultures grown with diffuse sunlight. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 5: Synechococcus cultures grown with diffuse sunlight. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 6: Cyanothece cultures grown with diffuse sunlight. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 7: Anabaena cultures grown with diffuse sunlight. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 8: Green Berries grown with white LED enrichment. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 9: Green Berries grown with 470nm LED enrichment. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 10: Green Berries grown with 521nm LED enrichment. Each of the tiles in this array represents a spectral-CSLM auto fluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 11: Green Berries grown with 521nm LED enrichment. Note blue-shifted fluorescence emission of the filaments in the upper right (Regions 11-13). See previous Figure legends for a description of the representation of the data in the figure. Figure 12: Green Berries grown with 595nm LED enrichment. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis). Figure 13: Green Berries grown with 380nm LED enrichment. Each of the tiles in this array represents a spectral-CSLM autofluorescence scan of a circular area (one cell) in the micrograph. The color indicates the intensity of the fluorescence emission (legend) and it's wavelength (y axis) detected at one of the seven laser excitation wavelengths (x axis).