**2. Fluorescence microscopic spectroscopy**

Modern fluorescence microscopic spectroscopy (FMS) or confocal laser scanning microscopic spectroscopy provides a unique opportunity to obtain high-resolution images and intrinsic fluorescence emission spectra from single cyanobacterial cells [42–45]. Moreover, using spectral unmixing, the fluorescence of individual spectral components can be resolved, and their relative intensities can be calculated [46–48]. Unfortunately, most of the researches use confocal laser scanning microscopy only for imaging [49–54]. In this part, the attention is paid mostly to the spectroscopic studies by means of CLSM, and we will give some guidelines on methods of investigation and sample preparation.

5–9. 1 pixel corresponds to 53.5 × 53.5 nm. The photomultiplier (PMT) voltages were used in range from 900 to 1100 V. The fluorescence emission images were accompanied with the transmission images (in the parallel channel), collected by a transmission detector with the photomultiplier voltages ranged from 300 to 500 V. For better signal yield, lambda scans were performed with "low speed" setting (400 Hz) in bidirectional scan mode and with a pinhole setting of 1 Airy unit (the inner light circle of the diffraction pattern of a point light source, corresponds to a diameter of 102.9 μm with the lens used (see [46]). Regions of interest (ROIs) representing single cells or subcellular regions were used to calculate fluorescence spectra.

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In CLSM applications, the laser light density in the focus point is high. But, generally, it is difficult to compare the excitation energies used in CLSM with those from methods developed to measure photosynthetic parameters. In CLSM, light is deposited in short "dwell times" during the laser scanning process. Dwell time and the intervals between the illuminations may influence photo-damage and saturation of photosynthesis. Thus, since most chromophores bleach under the high laser excitation energies, a bleach-test should be performed [43]. It was shown experimentally that especially phycoerythrin (PE) and phycocyanin (PC), as an accessory pigments, were very sensitive to photo-bleaching, while the fluorescence of Chlorophyll *a* (Chl *a*) and allophycocyanin (APC) remained stable in the intact cells [43]. During the detection, the fluorescence of the main accessory pigments for each cyanobacterial strain should be controlled and the changes in their fluorescence should not exceed 10–20%. The power of individual laser lines should be chosen according to the photodamage they cause. In our experiments, the repeated spectra were obtained under selected excitation power at a fixed point in a cell to check whether the excitation would affect the cells. It was shown that at the above chosen excitation energies, the fluorescence spectra did not vary within the experimental error during 10–15 records. When excitation energy was increased, both the height and the center of the bands varied enormously with time because of photodamage or structurebreakdown in photosynthetic systems. In the experiments, where several laser lines were involved for the investigation, the first spectrum was recorded again at the end of each series to control the initial state of the cell. To compare different cells in one physiological state, the fluorescence spectra were taken from the cells of one strain cultured at different days and it was established that the variations in spectrum shape and intensity are not considerable. To visualize differences between strains with higher spectral and spatial resolution, lambda scans were performed with 6 nm bandwidth and with 6 nm steps. As far as the fluorescence intensities depend on the excitation energy (which varies for different laser lines), sensitivity setting of the photomultiplier, and the distance from the sample, all spectra were usually

normalized to their maximum and only qualitative analysis was performed.

**2.3. A set of excitation wavelengths should be used during the investigation**

It is well known that phycobilisome contains several kinds of biliproteins, and its absorption and fluorescence spectra reflects the contribution of each. On the other hand, as a result of energy transfer among the tightly coupled biliproteins in the phycobilisome, fluorescence of the intact living cyanobacterial cells is originated from the efficiency of the energy transfer between these components and each transfer step appears in the spectrum shape as peak or

**2.2. Prevent photobleaching**

It is well known that measurement and analysis of fluorescence is one of the most powerful ways to probe photosynthetic systems because it reports on the energy transfer and trapping. This fluorescence originates from excited states that were lost before photochemistry took place. It usually represents a small fraction of the excited state decay in a functional photosynthetic complex. Nevertheless, this small fraction can be easily detected by CLSM. With the confocal fluorescence microscopy, a very small excitation and detection areas can be investigated, so that single cells under non-damage conditions can be studied *in vivo*. Although, the pigment structure of different cyanobacterial strains has been intensively investigated, the variations of *in-vivo* operation of photosynthetic apparatus for different cyanobacterial species have not been analyzed yet. We suppose that the best way to investigate the operation of photosynthetic system *in vivo* is a single-cell fluorescence spectroscopy. Single-cell detection can provide the information on small peculiarities that is regularly buried in normal ensemble average experiments. This is thus a good way to study the time evolution process and spectroscopic properties of individual cells. Both steady-state and time-resolved fluorescence measurements can be used for probing the organization and functioning of photosynthetic systems by means of CLSM.

#### **2.1. CLSM parameters**

In the presented investigation, Leica TCS-SP5 was used for spectral CLSM of living cyanobacterial cells. Fluorescence emission spectra of the intact cells were measured at eight excitation wavelengths corresponding to all available laser lines. The excitation wavelengths are: 458, 476, 488, 496, and 514 nm—the lines of Ar laser, 405 nm is the line of diode UV laser and 543 and 633 nm are the lines of HeNe laser. In the experiments, presented below in this chapter, laser power settings were as follows: 29% of Ar laser power was reflected onto sample with acousto-optical tunable filter (AOTF) and further power percentage for its laser lines was: 30% of 458 nm laser-line and 10% for all other lines. 405 nm line of diode UV laser was reflected onto sample with 3%, HeNe laser lines 543 and 633 nm were reflected with 10 and 2%, respectively. An acousto-optical beam splitter (AOBS) was used to transmit sample fluorescence to detector. Emission spectra between 520 and 785 nm were recorded using the lambda scan function of the "Leica Confocal Software" by sequentially acquiring a series ("stack") of 38–45 images, each with a 6-nm fluorescence detection bandwidth and with 6 nm wavelength step. For obtaining fluorescence-intensity information images of 512 × 512 pixels were collected with a 63× Glycerol immersion lens (glycerol 80% H<sup>2</sup> O) with a numeric aperture of 1.3 (objective HCX PL APO 63.0 × 1.30 GLYC 37°C UV) and with additional digital zoom factor 5–9. 1 pixel corresponds to 53.5 × 53.5 nm. The photomultiplier (PMT) voltages were used in range from 900 to 1100 V. The fluorescence emission images were accompanied with the transmission images (in the parallel channel), collected by a transmission detector with the photomultiplier voltages ranged from 300 to 500 V. For better signal yield, lambda scans were performed with "low speed" setting (400 Hz) in bidirectional scan mode and with a pinhole setting of 1 Airy unit (the inner light circle of the diffraction pattern of a point light source, corresponds to a diameter of 102.9 μm with the lens used (see [46]). Regions of interest (ROIs) representing single cells or subcellular regions were used to calculate fluorescence spectra.

### **2.2. Prevent photobleaching**

**2. Fluorescence microscopic spectroscopy**

16 Cyanobacteria

methods of investigation and sample preparation.

systems by means of CLSM.

**2.1. CLSM parameters**

Modern fluorescence microscopic spectroscopy (FMS) or confocal laser scanning microscopic spectroscopy provides a unique opportunity to obtain high-resolution images and intrinsic fluorescence emission spectra from single cyanobacterial cells [42–45]. Moreover, using spectral unmixing, the fluorescence of individual spectral components can be resolved, and their relative intensities can be calculated [46–48]. Unfortunately, most of the researches use confocal laser scanning microscopy only for imaging [49–54]. In this part, the attention is paid mostly to the spectroscopic studies by means of CLSM, and we will give some guidelines on

It is well known that measurement and analysis of fluorescence is one of the most powerful ways to probe photosynthetic systems because it reports on the energy transfer and trapping. This fluorescence originates from excited states that were lost before photochemistry took place. It usually represents a small fraction of the excited state decay in a functional photosynthetic complex. Nevertheless, this small fraction can be easily detected by CLSM. With the confocal fluorescence microscopy, a very small excitation and detection areas can be investigated, so that single cells under non-damage conditions can be studied *in vivo*. Although, the pigment structure of different cyanobacterial strains has been intensively investigated, the variations of *in-vivo* operation of photosynthetic apparatus for different cyanobacterial species have not been analyzed yet. We suppose that the best way to investigate the operation of photosynthetic system *in vivo* is a single-cell fluorescence spectroscopy. Single-cell detection can provide the information on small peculiarities that is regularly buried in normal ensemble average experiments. This is thus a good way to study the time evolution process and spectroscopic properties of individual cells. Both steady-state and time-resolved fluorescence measurements can be used for probing the organization and functioning of photosynthetic

In the presented investigation, Leica TCS-SP5 was used for spectral CLSM of living cyanobacterial cells. Fluorescence emission spectra of the intact cells were measured at eight excitation wavelengths corresponding to all available laser lines. The excitation wavelengths are: 458, 476, 488, 496, and 514 nm—the lines of Ar laser, 405 nm is the line of diode UV laser and 543 and 633 nm are the lines of HeNe laser. In the experiments, presented below in this chapter, laser power settings were as follows: 29% of Ar laser power was reflected onto sample with acousto-optical tunable filter (AOTF) and further power percentage for its laser lines was: 30% of 458 nm laser-line and 10% for all other lines. 405 nm line of diode UV laser was reflected onto sample with 3%, HeNe laser lines 543 and 633 nm were reflected with 10 and 2%, respectively. An acousto-optical beam splitter (AOBS) was used to transmit sample fluorescence to detector. Emission spectra between 520 and 785 nm were recorded using the lambda scan function of the "Leica Confocal Software" by sequentially acquiring a series ("stack") of 38–45 images, each with a 6-nm fluorescence detection bandwidth and with 6 nm wavelength step. For obtaining fluorescence-intensity information images of 512 × 512 pixels were col-

(objective HCX PL APO 63.0 × 1.30 GLYC 37°C UV) and with additional digital zoom factor

O) with a numeric aperture of 1.3

lected with a 63× Glycerol immersion lens (glycerol 80% H<sup>2</sup>

In CLSM applications, the laser light density in the focus point is high. But, generally, it is difficult to compare the excitation energies used in CLSM with those from methods developed to measure photosynthetic parameters. In CLSM, light is deposited in short "dwell times" during the laser scanning process. Dwell time and the intervals between the illuminations may influence photo-damage and saturation of photosynthesis. Thus, since most chromophores bleach under the high laser excitation energies, a bleach-test should be performed [43]. It was shown experimentally that especially phycoerythrin (PE) and phycocyanin (PC), as an accessory pigments, were very sensitive to photo-bleaching, while the fluorescence of Chlorophyll *a* (Chl *a*) and allophycocyanin (APC) remained stable in the intact cells [43]. During the detection, the fluorescence of the main accessory pigments for each cyanobacterial strain should be controlled and the changes in their fluorescence should not exceed 10–20%. The power of individual laser lines should be chosen according to the photodamage they cause. In our experiments, the repeated spectra were obtained under selected excitation power at a fixed point in a cell to check whether the excitation would affect the cells. It was shown that at the above chosen excitation energies, the fluorescence spectra did not vary within the experimental error during 10–15 records. When excitation energy was increased, both the height and the center of the bands varied enormously with time because of photodamage or structurebreakdown in photosynthetic systems. In the experiments, where several laser lines were involved for the investigation, the first spectrum was recorded again at the end of each series to control the initial state of the cell. To compare different cells in one physiological state, the fluorescence spectra were taken from the cells of one strain cultured at different days and it was established that the variations in spectrum shape and intensity are not considerable. To visualize differences between strains with higher spectral and spatial resolution, lambda scans were performed with 6 nm bandwidth and with 6 nm steps. As far as the fluorescence intensities depend on the excitation energy (which varies for different laser lines), sensitivity setting of the photomultiplier, and the distance from the sample, all spectra were usually normalized to their maximum and only qualitative analysis was performed.

#### **2.3. A set of excitation wavelengths should be used during the investigation**

It is well known that phycobilisome contains several kinds of biliproteins, and its absorption and fluorescence spectra reflects the contribution of each. On the other hand, as a result of energy transfer among the tightly coupled biliproteins in the phycobilisome, fluorescence of the intact living cyanobacterial cells is originated from the efficiency of the energy transfer between these components and each transfer step appears in the spectrum shape as peak or shoulder (**Figure 1**). Moreover, depending on the excitation wavelength, the room temperature fluorescence emission spectrum of intact cyanobacterial cells exhibits various extents of contribution of phycobilisome emission to the spectrum. If one exclusively excites Chl *a*, using a 458 nm line of an Ar laser, the emission spectrum by cyanobacterial cells shows no appreciable emission of PC or APC. In cyanobacteria, the 458 nm excitation is preferentially absorbed by photosystem I (PSI) that contains more Chl *a* than by photosystem II (PSII) and is stoichiometrically more abundant than PSII. However, because reaction center of PSI turns over faster than the PSII, it has lower fluorescence intensity than the PSII antenna. This is indicated by PSI emission band at 715 nm which is much weaker than the PSII emission band at 682 nm. The excitation by intermediate (blue and green) wavelengths (405, 488, and 496 nm) reveals fluorescent maxima of all photosynthetic pigments, as the light in this range is absorbed by all pigment-protein complexes almost in equal portions and fluorescence emits by all steps of energy transfer chain (**Figure 1**). The direct excitation of cells in the PC absorption region at 514 and 543 nm, results in emission spectrum with two main peaks at 580 and 656 nm, which are due to PE, PC and APC emission and for species that lack PE the emission accumulates mostly near 656 nm. The spectra of the 633 nm excitation (not shown here) directly give a prominent emission band at 656 nm, that originates from PC, omitting band at 580 nm, which cannot be excited by 633 nm, even for species that have PE. Other small emission bands, corresponding to fine pigment structure of antenna complex, are not resolved in the room temperature investigation. Comparative analysis of the series of fluorescence spectra for different cyanobacterial species and strains reveals visible variations in their shape (**Figure 1**). If the fluorescence spectra were taken from alive cells in normal physiological state, which are cultured in the same growth environmental conditions, then the interspecies variations in pigment/chl *a* ratios are more pronounced than variations within the individual species. Species/strains differentiation could be carried out on the base of fluorescence analysis.

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As it was pointed out, the single-cell fluorescence spectra depend not only on the selected strain, but also on the physiological state of the chosen cell. CLSM is the only method that can accurately diverse alive and semi-dead cells (**Figure 2**), and compared to other direct fluorescence measurements, records the fluorescence emission spectra from only active pigments in alive cells and does not acquire information from dissolved pigments, organic substances, and debris around. Considerable differences in shape and intensity of the fluorescence emission spectrum of cyanobacterial cells within one strain in normal and depressed physiological states allow to estimate the viability of the whole culture relying not only on the visual methods, but also on the accurate spectral analysis. In **Figure 2** (right panel), three typical fluorescence spectra for three different physiological states of one microcystis cell are presented. Spectrum I corresponds to the alive cell, spectrum II is the spectrum of cell in the depressed physiological state, and spectrum III is a spectrum of the dead cell. All spectra were excited by 488 nm laser line. Here the normalization to maximum intensity was not made, so that to illustrate the considerable difference in intensities between normal and depressed cells. On the left panel, corresponding transmission and fluorescent images of the cell under

**Figure 2.** CLSM images and single-cell fluorescence spectra (in relative units) for cyanobacterial strain *Microcystis CALU 398*, obtained at three different physiological state of the same cell: I—spectrum of the alive cell, II—spectrum of the cell in depressed physiological state, and III—spectrum of the dead cell. The white bar corresponds to 2.5 μm. Dashed lines

indicates fluorescence wavelengths of APC, PC and Chl *a* at 650, 660 and 682 nm, respectively.

**2.4. Investigation of physiological state of single cell**

**Figure 1.** The examples of CLSM images and normalized single-cell fluorescence spectra for four cyanobacterial strains. The white bar corresponds to 25 μm. Corresponding excitation laser lines are indicated in plot legend. Dashed lines indicate fluorescence wavelengths of PC and Chl *a* fluorescence at 656 and 682 nm, respectively.

mostly near 656 nm. The spectra of the 633 nm excitation (not shown here) directly give a prominent emission band at 656 nm, that originates from PC, omitting band at 580 nm, which cannot be excited by 633 nm, even for species that have PE. Other small emission bands, corresponding to fine pigment structure of antenna complex, are not resolved in the room temperature investigation. Comparative analysis of the series of fluorescence spectra for different cyanobacterial species and strains reveals visible variations in their shape (**Figure 1**). If the fluorescence spectra were taken from alive cells in normal physiological state, which are cultured in the same growth environmental conditions, then the interspecies variations in pigment/chl *a* ratios are more pronounced than variations within the individual species. Species/strains differentiation could be carried out on the base of fluorescence analysis.

#### **2.4. Investigation of physiological state of single cell**

**Figure 1.** The examples of CLSM images and normalized single-cell fluorescence spectra for four cyanobacterial strains. The white bar corresponds to 25 μm. Corresponding excitation laser lines are indicated in plot legend. Dashed lines

shoulder (**Figure 1**). Moreover, depending on the excitation wavelength, the room temperature fluorescence emission spectrum of intact cyanobacterial cells exhibits various extents of contribution of phycobilisome emission to the spectrum. If one exclusively excites Chl *a*, using a 458 nm line of an Ar laser, the emission spectrum by cyanobacterial cells shows no appreciable emission of PC or APC. In cyanobacteria, the 458 nm excitation is preferentially absorbed by photosystem I (PSI) that contains more Chl *a* than by photosystem II (PSII) and is stoichiometrically more abundant than PSII. However, because reaction center of PSI turns over faster than the PSII, it has lower fluorescence intensity than the PSII antenna. This is indicated by PSI emission band at 715 nm which is much weaker than the PSII emission band at 682 nm. The excitation by intermediate (blue and green) wavelengths (405, 488, and 496 nm) reveals fluorescent maxima of all photosynthetic pigments, as the light in this range is absorbed by all pigment-protein complexes almost in equal portions and fluorescence emits by all steps of energy transfer chain (**Figure 1**). The direct excitation of cells in the PC absorption region at 514 and 543 nm, results in emission spectrum with two main peaks at 580 and 656 nm, which are due to PE, PC and APC emission and for species that lack PE the emission accumulates

18 Cyanobacteria

indicate fluorescence wavelengths of PC and Chl *a* fluorescence at 656 and 682 nm, respectively.

As it was pointed out, the single-cell fluorescence spectra depend not only on the selected strain, but also on the physiological state of the chosen cell. CLSM is the only method that can accurately diverse alive and semi-dead cells (**Figure 2**), and compared to other direct fluorescence measurements, records the fluorescence emission spectra from only active pigments in alive cells and does not acquire information from dissolved pigments, organic substances, and debris around. Considerable differences in shape and intensity of the fluorescence emission spectrum of cyanobacterial cells within one strain in normal and depressed physiological states allow to estimate the viability of the whole culture relying not only on the visual methods, but also on the accurate spectral analysis. In **Figure 2** (right panel), three typical fluorescence spectra for three different physiological states of one microcystis cell are presented. Spectrum I corresponds to the alive cell, spectrum II is the spectrum of cell in the depressed physiological state, and spectrum III is a spectrum of the dead cell. All spectra were excited by 488 nm laser line. Here the normalization to maximum intensity was not made, so that to illustrate the considerable difference in intensities between normal and depressed cells. On the left panel, corresponding transmission and fluorescent images of the cell under

**Figure 2.** CLSM images and single-cell fluorescence spectra (in relative units) for cyanobacterial strain *Microcystis CALU 398*, obtained at three different physiological state of the same cell: I—spectrum of the alive cell, II—spectrum of the cell in depressed physiological state, and III—spectrum of the dead cell. The white bar corresponds to 2.5 μm. Dashed lines indicates fluorescence wavelengths of APC, PC and Chl *a* at 650, 660 and 682 nm, respectively.

investigation are shown. Analyzing the shape of the ensemble average fluorescence spectrum and counting the relative number of alive and semi-dead cells the conclusion about the viability of the whole culture at given developmental stage could be made.

and technological study. In addition, the results of single-cell spectroscopic analysis are much more suitable for further statistical and analytical calculations then the conventional optical methods of investigations. In this chapter, we present several examples of practical applica-

Fluorescence Microscopic Spectroscopy for Investigation and Monitoring of Biological Diversity…

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21

Since the first broad-scale spectroscopic investigations, the authors of many articles note the dependence of the intrinsic fluorescence spectra of cyanobacteria on the developmental stage of the culture and physiological state of single cells. It is well-known that the light-harvesting and energy-transfer capacities of phycobilisomes can react to the environmental changes, as well as to the changes in physiological state of the living cells induced by stress conditions [39, 64, 73–75]. However, this effect has not yet been widely used to assess the viability of the culture. Several authors pointed out that, although a single-cell fluorescence spectra for the diverse physiological states differ significantly, the physiological state of the given cell cannot be estimated correctly because of the absence of a full set of reference spectra [40, 76, 77]. Moreover, the authors of [40, 76, 77] pointed out that while comparing spectra of individual cells and the results of the ensemble average experiments at a culture as a whole (so-called integral spectra), a significant difference was observed. Obviously, this difference is owing to a wide diversity of single-cell physiological states in the bulk growing culture, which in sum gives different integral fluorescence spectra for a specified strain at different developmental stages because of the variations in cell's proportions. This, of course, should be taken into account. On the other hand, the whole culture in addition to a set of single living cyanobacterial cells consists of metabolites, dissolved pigments, other organic substances and cellular debris. All these substances form undesirable

Actually, the intensity of fluorescence emitted by single photosynthetic cell *in vivo* depends only on the structure and operational effectiveness of photosynthetic apparatus, tracing intime physiological state of the cyanobacterial cell. Thus, the fluorescence emission can be used effectively to monitor various physiological processes. **Figure 3** illustrates the temporal changes of in-vivo fluorescence spectrum taking place in one living cell of cyanobacteria strain *Synechocystis CALU 1336* under light and heat stress. On the other hand, this timeline set of fluorescence emission spectra illustrates all stages of cyanobacterial cell degradation, that is, all possible physiological states. It is obvious that during the evolution of the culture and aging of each cell all this stages will be presented in the natural samples simultaneously.

Several newer publications [76–80] clearly demonstrate that the variations in the fluorescence shape and intensity of living cells, presented in **Figure 3**, indicate the consequent degradation in the light harvesting chain (antenna complex—reaction center) and following dissociation of the detached antenna complex. It can be seen that in the alive cell, the chlorophyll *a* fluorescence prevails over the fluorescence of the pigment-protein complexes of phycobilisome. While the single-cell physiological state changes for the worse, the photosynthetic apparatus shows instability in operation, that is, the most part of the absorbed energy emits as fluorescence at the early stages of light harvesting. At the last stages, the changes in fluorescent spectrum, shown in **Figure 3**, are the same as it was demonstrated in the works [80–82], where

**3.1. Monitoring of physiological state of single cyanobacterial cell and a whole** 

and unpredictable fluorescent background in volume samples.

tion of the described CLSM technique.

**culture**

### **2.5. Sample preparation**

Special attention should be paid to the sample preparation, as well as we work with alive objects. Coverslip should be pressed very carefully to prevent any glass slide, which can cause cell damage. On the other hand, one should keep in mind that cyanobacteria can move and glide; so to fix the object, the coverslip should be pressed hard enough to prevent any motility of the investigated object, which sometimes have a diameter near 1 μm (e.g., microcystis and synechocystis cells).

#### **2.6. Spectral unmixing**

There is another very powerful tool implemented in CLSM—spectral unmixing. Unfortunately, in living cyanobacterial cells, it meets some difficulties. The authors of [43] pointed out that a lot of problems arise during the spectral unmixing procedure, which is based on the spectra of isolated phycobiliproteins. These problems are caused by the fact that the light absorption and emission properties of isolated phycobiliproteins are rather different from those of the intact phycobilisomes in the living cyanobacterial cells. In living cells, the spectral properties of pigments from certain organisms may differ crucially from the properties of the dissolved ones, for example, spectra of the components can vary in peak widths and may be shifted in wavelength due to different pigment-protein and linker connections. Thus, the analysis based on the initial fluorescence spectrum without any decomposition is preferable for living cells.
