STP/RSTP

The discovery of a green fluorescent protein (GFP), which was isolated from the jellyfish Aequorea victoria (Shimomura 1962), along with its gene fusion (C. Prashera et al. 1992; Tsien 1998) has revolutionized the field of cell biology. Due to their versatility, specificity and quantitative capabilities for the live-cell imaging, fluorescent proteins fused to their target proteins have become crucial tools in cell biology experiments and have allowed the emergence of single-cell studies of gene expression.

Molecular cloning methods of fusing the fluorophore moiety to a protein of interest (e.g. an enzyme or an RNA polymerase subunit), have helped researchers to monitor cellular processes in living systems. Due to rapid evolution, fluorescent proteins can now cover the visible spectral wavelengths. Also enhanced were properties such as maturation and degradation times, folding, oligomerization, brightness, and photostability. Also, these improvements have helped to perform multicolor imaging of any protein (Shaner et al. 2004) and allow us to study the subcellular architecture at the sub-nano second-time resolution (Tsien 1998). Overall, fluorescent proteins have become essential tools in areas ranging from studies of the complex behavior of single-molecules, of the internal dynamics of the molecular process, quantitative studies of gene expression dynamics, etc. (Yu et al. 2006; Stracy & Kapanidis 2017; Golding et al. 2005).

During the last two decades, this field also has produced advanced fluorescent probes with unique characteristics, such as photoactivation and photoconversion (Daya & Davidson 2009; Wu et al. 2011). These modified fluorescent proteins can be switched on and off or, be converted from the non-fluorescent state to the stable fluorescent state in response to light at an appropriate wavelength. These works have motivated many advances in microscopy imaging techniques, e.g., super-resolution microscopy, that enables the monitoring of inner components of cells in great detail.

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Some factors have to be carefully considered when conducting imaging experiments. For example, the brightness of fluorescent proteins should be above the cellular background level, their photostability needs to be robust, and there should be minimum cross-talk between the emission and excitation spectra. Also, when fused with the target protein, the effect of the fluorescent protein on the native protein functionality should be as weak as possible (Shaner et al. 2004).

Although fluorescent proteins have several advantages, drawbacks of using them include fluctuations in the fluorescent intensity (Ha & Tinnefeld 2012) due to changes in the environmental conditions and photostability. For example, some wild-type GFP is sensitive to temperature, while Yellow fluorescent proteins (YFP) are sensitive to pH and chloride (Wachter & Remington 1999). To overcome this situation, several altered or mutated versions of fluorescent proteins (Shaner et al. 2004) ( see Figure 3.1) has been developed, in order to improve stability, folding and sensitivity to environmental conditions.

Figure 3.1: Purified fluorescent proteins shown in visible light. These fluorescent proteins from left to right (mHoneydew, mBanana,

mOrange, tdTomato, mTangerine, mStrawberry, and mCherry) were derived from the Discosoma sp. red fluores- cent protein. This image is extracted from (Shanner et al. 2004) with permission from the Nature publishing group.

In addition, when performing live-cell imaging, one common goal is to monitor cellular dynamics, which requires fast image acquisition. This demands short exposure times, meaning that fluorescent proteins that absorb and emit light have to be significantly bright than the cellular background (Ha & Tinnefeld 2012). This requests a highly sensitive detector and a bright light source. Further, the numerical aperture needs to pay attention when choosing the objective.

The most common optic system used in the illumination of fluorescence microscopy is a wide-field epi- illumination. In this optic system, the entire area of the sample is exposed to light either from above (in a standard upright microscope) or from below (in an inverted configuration). It excites the incident lamp excitation light of an area of ׽10 x 10 —m2 _{(Webb & Brown 2012). Thus, the volume illuminated is quite}

large, causing out-of-focus fluorescent molecules to contribute to the background fluorescence signal. Wide-field epi-illumination has a wide range of applications in bacterial studies, such as monitoring the dynamics of fluorescently tagged RNA molecules (Golding et al. 2005) and protein molecules (Yu et al. 2006). On the other hand, there are several limitations such as low resolution, excess of out of focus fluorescent signal and, photobleaching of the sample.

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Figure 3.2: Lightpath illustration of wide-field epi-illumination and confocal microscopy. In confocal microscopy (B), light from the

light source (Ls) is focused through a pinhole for illumination (Pi) and subsequently passes through a sample (S), resulting in a small focal volume. In wide-field epifluorescence microscopy (A), the entire sample volume is exposed to light.

To avoid such limitation, several other methods have been developed, including Confocal microscopy (Pawley 2006), Total Internal Reflection (TIRF) microscopy (Fish 2015), and Highly Inclined and Laminated Optical (HILO) sheet microscopy (Tokunaga et al. 2008). The primary goal of this microscopy is to eliminate excess of out of focus fluorescent light during the imaging process by restricting the illumination volume of the sample.

In confocal laser scanning microscopy, the light source for exciting the fluorescence molecule comes from the laser unit, and it is targeted to a region of interest. It can be used to obtain optical sections through a sample to exclude out of focus and background fluorescence (Figure 3.2). This is achieved with a pinhole aperture where excited light passes through a focal volume of a sample (Pawley 2006). The drawback of this optical system is the slowness of point scanning image acquisition, which restricts the area of the image. This speed can be increased by using spinning disc confocal microscopy that illuminates multiple regions of the sample and minimizes photobleaching or phototoxicity (Frigault et al. 2009; Nakano 2002). When comparing TIRF microscopy with confocal microscopy, a better optical section of the sample is illuminated. TIRF uses an evanescent wave, which is generated when the incident light is reflected at the interface of two transparent media with different refractive indices. TIRF allows to selectively illuminate and excite the fluorophores in a restricted region of the sample. As the energy of the evanescent wave field decreases exponentially with distance from the interface, the only fluorophore at a certain distance from coverslip is excited, which allows creating images with an outstanding signal-to-noise ratio. Also, TIRF can illuminate the region of the sample with an outstandingly high axial resolution, below 100nm. As a

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result, it can only probe molecules close to coverglass surfaces, e.g., membrane-associated molecules. To illuminate the region deeper than the TIRF range, HILO microscopy was developed (Tokunaga et al. 2008). HILO is generated by intense laser illumination, angled through a high numerical aperture objective to a sample, resulting in lower out-of-focus light signal (Tokunaga et al. 2008).

The methodologies above allow monitoring of fluorescent molecules in-vivo and in-vitro. Some of our studies also require the visualization of high contrast images of transparent live cells. Such images were acquired by phase-contrast microscopy. It employs an optical mechanism that converts minute differences in a phase into corresponding variations in amplitude, which can be seen as a difference in image contrast (Zernike F 1942). Phase-contrast microscopy enables to examine live cells, without exposing them to laser or staining dyes. It is one of the few methods available to quantify cell structure, shape, and size.